Beyond Extreme Close-Up Photography 9781785004667

Table of contents :
Cover Page
Title Page
Copyright Page
Contents
Preface
1 What is Magnification?
2 What Cameras are Best?
3 Techniques to go Beyond Macro
4 An Introduction to Microscopes
5 Lighting Extreme Close-ups
6 Support and Preparation
7 Focus Stacking
8 Inspiration for Extreme Close-up Photos
References and Further Reading
Index

Citation preview

Beyond Extreme Close-Up Photography

Beyond Extreme Close-Up Photography

Julian Cremona

CROWOOD

First published in 2018 by The Crowood Press Ltd Ramsbury, Marlborough Wiltshire SN8 2HR www.crowood.com This e-book first published in 2018 © Julian Cremona 2018 All rights reserved. This e-book is copyright material and must not be copied, reproduced, transferred, distributed, leased, licensed or publicly performed or used in any way except as specifically permitted in writing by the publishers, as allowed under the terms and conditions under which it was purchased or as strictly permitted by applicable copyright law. Any unauthorised distribution or use of thistext may be a direct infringement of the author’s and publisher’s rights, and those responsible may be liable in law accordingly. British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library. ISBN 978 1 78500 466 7

Cover Photograph: Eye of the small elephant hawkmoth. Forty images have been focus stacked to create this composite image using Helicon Focus software. (×6 magnification, Canon 7D mk2, 65mm MPE lens ƒ5.6 ISO 100, with diffuse twin macro flash.) through several rinses in clean water to remove detritus. Frontispiece: Part of the underside of a rose chafer beetle, Cetonia aurata. (×7 magnification, Canon 7D mk2 with 65mm MPE, 32mm of extension tube, ƒ5.6 ISO 100 twin macro flash, composite of thirty-two images.)

CONTENTS

Preface

1 What is Magnification? 2 What Cameras are Best? 3 Techniques to go Beyond Macro 4 An Introduction to Microscopes 5 Lighting Extreme Close-ups 6 Support and Preparation 7 Focus Stacking 8 Inspiration for Extreme Close-up Photos References and Further Reading Index

Preface

‘The contribution of visible life to biodiversity is very small indeed.’ Sean Nee writing in Nature, 2004. am a regular reader of the photographic press. The articles I enthusiastically lap up are as diverse as possible because this is one of my pet loves; taking photographs. My grandchildren know, just like their parents, that a camera will always be there to record the moment. Any moment of their lives. One, which some find strange, is the appearance and disappearance of teeth as they grow up; these can present technical challenges along with the personal level of dealing with my model. This should not come as a surprise to people who know me, because they realize that my passion is for anything natural and biological. The primary purpose of my travels around the globe is in the search for wilderness and then to record, through my photography, the life that exists there no matter what the size or type of organism. Learning through this form of research, I am better able to understand the dynamics of the natural world and go on to help in conservation and education.

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Spanish scorpion photographed at night with ‘black light’ (365nm wavelength), invisible to human eyes. In daylight the normal colour is yellow. Although this image is not an extreme close-up, it demonstrates that by photographing animals and plants in UV or infrared, some species become more striking and visible to us. This leads me to a conundrum. I have met people who just photograph flowers (waiting for the insects to disappear first). There is the bird photographer: I was in a very small boat on a river in a remote part of Costa Rica with a guide. I was sharing this with a German photographer, tripod and very long lens at the ready. An exotic bird appeared and we both got snapping. A howler monkey in the tree above;

crocodile up close baring its teeth in the water; a bright pink dragonfly at rest on the reed; only my camera was in action. The bird photographer would hardly look. Now back to the photographic magazines. I look at the wonderful wildlife photos. My interpretation of wildlife seems to be very different to others. Mammals like foxes, badgers, deer, elephants and tigers grace the pages along with birds and flowers. But surely flowers are not wildlife, are they? Or insects and sea anemones in a tide pool? Of course they are! I, and most biologists, fail to categorize creatures like the press because to me, all life that is wild should include all nature without exception. And here we reach the real ambiguity, because the vast majority of life on Earth is not visible to the naked eye. The photos we typically see in magazines, over and over, are just the tip of a huge biodiversity iceberg and 99.9 per cent of wildlife photographers are only taking photos of that tip. The enormity of what is left is largely overlooked. Some people will discover more as they use a macro lens to take photos of butterflies and other insects (even macro is seen as a different genre to wildlife in many magazines). There are a host of books and websites out there describing macro techniques, including ‘extreme macro’, but I am amazed that the sites are full of bees and flies. Where are the rest, such as beetle life in a pond or crabs on the seashore? There is so much more out there and still I am talking about the visible creatures, because almost all the biodiversity of life on our planet is not visible to our naked eye. Much of this has never been photographed and it will be in your garden, parkland or nearest wood or pond. Your range of subjects is vast and in practical terms inexhaustible.

The marine pseudoscorpion, Neobisium, photographed in 1998 with a Fuji bridge camera and an SLR lens reversed on to the front and coupled. (×6 magnification, live specimen in seawater, natural light.) Twenty years ago I photographed a tiny creature called a marine pseudoscorpion on the rocky shores of west Wales. A close relative of the true scorpion, it is a few millimetres long and while being quite common, is rarely seen because of its size. I struggled to find it but on this occasion, I took a photo on a Fuji bridge camera and for the ×4 or ×5 magnification, I had a 50mm SLR lens reversed over the front (a technique called coupling). Until very recently if you Googled this little arachnid, only my rather soft photo appeared as I had it on an educational website. What this illustrates is that we are passing a vast array of species every second of the day, but

very few photographers are taking advantage of that bonanza. Instead, like landscape photographers travelling to the classic honeypot sites, there are the nature photographers imaging the ‘big stuff’. I am not complaining as I do the same. Part of my life-long career in biology has been to educate children across the world in conservation issues. Once I stood up in a Singaporean auditorium and talked to a thousand teenagers, asking them, ‘What is the point of a panda? Given the choice, would you conserve the panda or a bluebottle fly?’, daring them to understand that if we lost pandas the world would carry on; a very sad and genetically poor world, but one that would survive. You can imagine the uproar. Flies are not cuddly creatures and so do not tug at the conservation heartstrings. We need flies to clear up our world of the dead and decaying, not to mention the mountains of dung produced. For the sake of conservation I believe we should be highlighting the wildlife that is so hidden. Entering the realm of the micro-world is such a cliché but it is also very true that it is an exciting, alternative world which so few people ever get to see, let alone photograph. The problem is down to the difficulty of taking photos of very small things and I have tried here and in previous books to redress this problem. To that end, in 2014 Crowood Press published my first book in the ‘Art and Techniques’ series, Extreme Close-up Photography and Focus Stacking. Discussing everything macro, it covered the area between around a fifth life-size (1:5) down to ×5 magnification (5:1). In 2017, Crowood published An Introduction to Digital Photomicrography by Brian Matsumoto and Carol Roullard. In contrast, this book covers photography of nature with magnifications well in excess of ×100, where the microscope is really the only tool available. I am writing this book to fit between these two and inevitably there will be a small degree of overlap. Starting beyond macro, we are aiming at the rather difficult ×5 (5:1) to ×30 (30:1) magnification region, although the book will in effect be covering anything between life size up to ×100. Some of the techniques will unavoidably have been mentioned in my other book but I will be taking these forward into new ideas. Most importantly, low cost and DIY systems will be emphasized as much as possible. Some people like categories and names and plenty are bandied about, often incorrectly, like macro photography – which is life size, 1:1. True photomacrography is the name we give to employing optics that are designed for use without an eyepiece on extension tubes or bellows. This can produce the magnifications of up to 40:1 or higher. Photomicrography is the term reserved for photography that requires a compound microscope using an eyepiece connected through a tube to an objective lens at the other end, where the subject is located. The magnifications will overlap, as it can start around 10:1, rising to 2,000:1 (considerably outside the realms of this book). In fact, the first chapter tackles the very issue of what we mean by magnification. Subsequent chapters consider cameras, techniques and equipment, lighting, supporting methods, focus stacking and at the end, a series of suggestions to try and develop your extreme close-up ideas. In particular, that last chapter helps consolidate

some of the detail of the book by bringing the theory into practice with hopefully inspiration on subjects you might never have considered.

While there are only around 500 species of marine fungi across the oceans, these tiny organisms are vital for recycling. The strands of a seashore fungus are growing on a red seaweed, possibly the first photograph ever taken of this species but as so little is known about them, even identification is difficult. (×8 magnification, composite of twenty images taken in seawater.) I run workshops and during this time, I seem to have developed a number of nicknames based on my extensive use of Blu Tack (other varieties are available, like white tack) and gaffer tape for holding everything together. You can never have enough! Over the years I am sure the people I have met on these workshops and at Quekett Microscopical Club meetings have taught me much more than I have taught them. Two from the club have allowed me to use a few of their amazing photos in this book, namely Mike Crutchley (who, living nearby, has many of his ideas borrowed and used in this book) and Mark Papp. The inspiration from being a member of the Quekett is quite overwhelming and I must thank Phil Greaves, Chris Ramon, Carel

Sartory, Ray Sloss, David Spears and Spike Walker for their invaluable help over the years. Phil has been good enough to read through a draft of the book and I thank him for his comments, but the mistakes are entirely my own. My wife Brenda is always there with support and ideas – thank you. Remember, there are more micro subjects waiting to be photographed in the square mile around your house than you can probably photograph in your lifetime. You just need a hand lens and time to stop and look carefully. Then attempt a photograph and the chances are you are the only person to have done that.

Chapter 1

What is Magnification?

he answer to this question may seem obvious and not that many years ago, it was even more simple to respond. Sherlock Holmes would have reached in his pocket and retrieved a large magnifying glass and the object would double in size to make it clearer to see. But today with a plethora of digital devices, the explanation has become distorted and more difficult to interpret.

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Fig. 1.1 Head of an opossum shrimp, photographed live in seawater with dark field lighting. Common in estuaries and on the shore, the family of mysids to which it belongs are believed to be a primitive group of crustaceans. (×4 magnification, Canon 7D mk2 with 65mm MPE lens at ƒ5.6, composite of 25-image stack.)

WHAT DO WE MEAN BY MAGNIFICATION?

With so much nutritional information needing to be put on food labels these days, manufacturers will typically make the font size small, sometimes so small it is almost impossible to read. As we age, the ability to resolve detail with our eyes can deteriorate, particularly in low light. Perhaps it is a feature of our modern age that legal information at the bottom of documents is in a tiny font, so we ignore it. Few people carry a large magnifying glass but invariably a smartphone will be in the pocket or bag. A quick snapshot of the label or document appears on the screen and with a pinch or two of your fingers, the image is magnified so the text can now be read. Clearly, this is one way in which we can magnify a subject. With a large enough screen, we can blow up our photos so that an image of our dog is so enlarged we might make out a tick on the side of the body. We could crop the image so that we just have the small part we want and then on the screen, it appears that we have a highly magnified subject. To produce anything vaguely useful, this technique relies on a high number of pixels being present in the original photograph as cropping the photograph possibly removes many millions of pixels.

Fig. 1.2 Green hairstreak butterfly photographed on a gorse flower. The photo is true macro 1:1 as the width of the image measures 35mm and in real life, the subject matter was 35mm across. The green hairstreak butterfly in Fig. 1.2 is an example. The image is an 18megapixel photo and to enlarge the head, this was cropped with the resulting photo becoming just 709 by 476 pixels, which is 337,484 pixels in total, making a 0.33 megapixel image. With so few pixels in the photo, it would appear very small on a screen and this can be improved by boosting the number, in this case to 2,500 by 1,678, around 4 megapixels. This can be done in any photo editor, followed by further enhancement to reduce any noise and increasing clarity and sharpening. It is printed small on this page to show that despite a reduction in quality, it may be acceptable.

Fig. 1.3 A crop of just the head of the green hairstreak in Fig. 1.2, showing it considerably enlarged but the image is of a significantly lower quality. The head of the butterfly may be magnified, but by how much? The original photograph covers an area approximately 35mm across and we could print a scale to indicate the size of the butterfly. This is usual in scientific literature but not always appropriate elsewhere. The cropped image is 3.5mm across and so if the images are printed to the same size, the enlarged section has been magnified by ×10. But this is

still rather vague and cropping is not the best way to magnify a subject and should be used when no other technique is available. If we wish to give a specific figure to magnification there has to be a better way. The conventional method harks back to film, in particular 35mm film. The green hairstreak butterfly would be classified as a life-size image or macro photograph. If the photo had been made on film, after processing, the size of the actual butterfly would be identical to the outline of the image on the film. You could lay one on top of the other and they would be the same. The field of view of the original is 35mm across, the same as in film. This true macro is given the ratio of 1:1, referring to the size on the film to the size of the subject in real life. If the photographer of this butterfly could have managed to get closer to the insect and fill the frame so that the back of the wing was touching the left side of the frame and the front leg was on the right frame edge (approximately 18mm across), the butterfly has now doubled in size and the ratio would be 2:1 as the image is now ×2; twice life size. The image could be enlarged on the screen or cropped to create what appears as a higher magnification but for this book, as is generally accepted, we will use the original photograph ratio as a measurement of magnification. If your photography is hovering around macro using a typical macro or close-up lens then working out the ratio will not be difficult. In fact most of the commonly available lenses will have these marked on the side of the lens barrel, even closefocusing zoom lenses. However, this book aims at significantly higher magnification and for any hope of accuracy in measuring this, you will need to photograph a sharply defined millimetre scale. A standard ruler will not do and unfortunately the desired scale can be quite expensive to buy. What is required is a glass microscope slide, etched with calibration down to 0.01mm. They are sold as a microscope stage micrometer scale and widely available. The stage refers to the fact that it is a glass slide to place on the microscope stage, not to be confused with a graticule that goes inside the eyepiece of a microscope for direct measurement of subjects as you look through the eyepiece at the specimen. Graticules are appreciably more expensive as well.

Fig. 1.4 Stage micrometer slide photographed to show a centimetre scale divided up so that the smallest divisions are 0.1mm (100 microns) across.

Fig. 1.5 Eye of a solitary bee and part of the antenna, photographed with Canon 7D mk2 fitted with bellows and objective lens.

Fig. 1.6 A reference photo of a stage micrometer scale is placed along the edge of the bee image, using the same equipment and set-up as the photo in Fig. 1.5 so that the magnification can be obtained. The red bar represents 0.1mm and can be added as a scale in scientific publications, but has not been included in subsequent photos in this book. Stage micrometers vary, but the most useful is the glass slide with three separate scales: 1mm, 0.1mm and 0.01mm. The latter is for higher microscope magnification and probably of least use for us here. When you are working with the different methods outlined in the book, for example, photographing the eye of a fly, as well as taking the insect, make sure you have created a reference photo. This reference is of the micrometer scale at that magnification. This only needs to be done once, labelled with the technique and kept safely in a separate folder. Then, if a photo is taken using

a particular set-up and you want the magnification, just match up the reference along the edge of the photo, as in Fig. 1.6. In the case of the bee’s eye a measurement of the length could be made (1.6mm). To calculate the magnification based on the photo:subject ratio, 35mm is divided by the reference scale of 1.73mm, approximately 20:1, which can be represented as ×20.

THE SIGNIFICANCE OF CAMERA SENSORS As we will see in the next chapter, the digital cameras vary considerably from one to another. There are only a few manufacturers of the sensors and camera producers generate different software (called ‘firmware’) for their cameras, which controls and determines the quality of the output. Unless you intend to crop heavily, the number of sensor pixels is less important than how big it is. Small sensors with large numbers of photosites (the areas that receive light and create the pixels) will be so tightly packed that they can cause interference that manifests itself as noise on the image: distracting random coloured pixels. Any size sensor can be used to produce high magnification, as we will see, and small sensors produce more depth of field (areas of detail that appear in focus).Sensor size seems to influence the magnification although this can be a rather abstract concept. Small sensors, like those in tiny compacts, can create what appears to be substantial magnification. Our discussion of magnification so far is related to 35mm film. A digital sensor, this size is found in a full frame camera and when lenses – such as a 100mm macro – are fitted to the body the focal length applies, that is, it will be 100mm. APS-C sensors are slightly smaller and the 35mm equivalent for the lenses will not be the same. Owners of Nikon and Sony cameras will be aware that they have to multiply by a crop factor of ×1.5 to identify the correct focal length while Canon APS-C users correct with a factor of ×1.6. The 100mm macro now becomes 150mm and 160mm, respectively. What is usually forgotten in this mental conversion is the value of the APS-C to magnification. With the macro set at 1:1, try photographing a millimetre scale. The full frame model will have a width of 35mm as expected (life size) but the APS-C models will be much less: between 22–24mm. Depending on the camera, divide 35mm by the crop factor, for example, Canon with ×1.6 is 21.9mm.

Fig. 1.7 Donacia beetle. A single image taken at ×7 magnification shows a limited amount of visible detail or depth of field.

Fig. 1.8 Donacia beetle. This is a composite of thirty-five images combined to create a huge depth of field. This is the process of focus stacking.

Fig. 1.9 Ruby-tailed wasp. This dead specimen was taken with a small (6.16×4.62mm) sensor Olympus TG-5 compact camera. As long as the image is not excessively enlarged, the sensor can produce a good image. (×3 magnification, LED light, autostacked image.) Or looking at this another way, a Canon macro photo taken with a macro lens on a 1:1 setting actually provides a magnification of ×1.6. For Nikon and Sony this will be ×1.5 and Pentax is ×1.53. APS-C sensors subtly increase magnification and this is important for what we discuss from here onwards. Additionally there will be a very slight improvement in depth of field. On the theoretical side, however, we see that sensors smaller than full frame are arguably not really increasing magnification, as all they do is crop the image. We seem to have gone full circle back to where we started! Not really, but it is worth bearing all this in mind. For the theoretically minded, we can work out the actual magnification of the sensor in any camera, especially a compact with a fixed lens. Firstly, find out the size of the sensor. This is very easily done online. For example, the Olympus TG-5 has a 1 × 2.3in or 6.16 × 4.62mm sensor. Set the camera on the closest focus it can achieve and photograph a ruler, with a result from the TG-5 of

around 19mm across the width of the shot. Divide the sensor width by this photographed width, 6.16 divided by 19, which gives 0.32. This is the true magnification of the camera or given as a ratio, 1:0.32; so not a true macro even though it has achieved a photograph which is similar to that in a full-frame camera. TABLE OF MAGNIFICATION AND FIELD OF VIEW A very useful tool is a magnification chart. As you develop your range of techniques, use the method above of photographing a scale so that a record can be kept of the specific magnification achieved. I do this in a spreadsheet that is constantly evolving as I try new ideas. The example below is just a clip from part of the spreadsheet. I regularly print this out and scribble over it with amendments and comments.

APS-C Mag

Methods

Full Frame Mag

Working Dist mm

Stack mm across Field Interval mm of View

Canon 65mm MPE ×1

1.6

1

95

0.4

22

×2

3.1

2

65

0.26

11.3

×3

4.7

3

50

0.16

7.45

×4

6.3

4

44

0.12

5.6

×5

7.9

5

40

0.09

4.45

10.0

6

38

0.05

3.5

13.5

8

7

0.03

2.6

19.4

12

6

0.02

1.8

6.7

4

0.1

5.2

100mm extension

10.1

6

0.05

3.45

200mm extension

14.6

9

0.02

2.4

×5 + 60mm extension Extension tubes with ×10 objective 13mm extension adapter

+

44mm extension adapter

+

Bellows objective

with

No extension

×4

PROBLEMS OF MAGNIFICATION Apart from all the theory behind magnification, the sheer joy of enlarging subjects and seeing an incredible world that you would otherwise be unable to appreciate is the excitement of extreme close-up photography. The more you magnify, the further you depart from what you knew and what is now completely alien. Unfortunately it comes with a few hurdles that have to be crossed and these will be covered within the

following chapters. It would be brilliant if we could learn and understand all the answers at once. As this is impossible, the key problems are being flagged up now so you are aware of them from the start and hopefully as the book progresses, the answers will be revealed.

Loss of Depth of Field (DoF) Depth of field, the amount of visible detail in a photo, varies with different lens apertures. Most noticeable is the loss of DoF with magnification so that at ×5 magnification at ƒ5.6, it can be 0.06mm and at ƒ16 not much better at 0.09mm. MANUAL OR AUTOFOCUS? Autofocus can be used to photograph a subject above 1:1 life size, although even at 1:4 it may be better to switch this off and manual focus to stop the camera ‘hunting’ back and forth. For closefocus and macro work, such as a butterfly, with autofocus switched off, set the lens focus at the approximate magnification required. Then focus on the subject by moving your body and camera (hold your elbows tight to your body) back and forth slowly until it is sharp in the viewfinder. I have been asked numerous times how I can see a small insect, raise the camera to my eye and within seconds have taken the photo. Many people lose the insect between seeing it and raising the camera. The trick is to keep both eyes open. Most people have a tendency to shut the eye that is not up at the viewfinder. I keep my left eye watching the insect as I raise the camera to my right eye and match the view as I begin moving in for the shot. The moment it is in focus, I fire the shutter. The camera is supported in the palm of my left hand with the right taking the photo. If the insect remains still I might then adjust the focus closer, using the fingers of the left hand and move in for an even closer photo. Practice on non-important material. Beyond 1:1, autofocus should be permanently switched off and in fact, the lenses and techniques discussed in Chapter 3 will largely be non-focusing ones. That means they all require the camera to be moved for focusing.

Loss of Light Even if the surrounding ambient light does not change, the closer to the subject you become, the less light is available. Also many of the methods we use reduce light. Photographing small objects, closer and closer, the available light becomes less of an option and the manipulation of the light will in some way be necessary.

Difficulty in Focusing and Composition Low light levels make it difficult to see when the subject is in focus. Using live view on a camera will help considerably and this will have a facility to magnify the live picture up to ×10, so that individual pixels can be seen. This is presuming that you can see the subject in the first place. Photographing a relatively flat specimen, for example, scales on the wing of a butterfly, is straightforward as while the majority is out of focus, you can still get a feel for everything that will finally make up the image. Above ×5–×10 magnification, three-dimensional subjects are difficult not only to focus but to

appreciate composition. Consider the two images in Fig. 1.10. To obtain a photograph of the entire head required a collection of around 200 images, which were then focus stacked (explained in Chapter 7). When trying to focus on the tip of the mouthparts, it is impossible to see where any of the rest of the head is located, just a vague smudge in the background. This makes positioning of the subject and camera quite difficult as so little is in focus. In the case of the harvestman, more than six different focus-stacked composites were made. After each stack the composite was prepared on the computer, checked and adjustments made to the composition of the specimen before starting another stack.

Fig. 1.10 Focusing at ×12 magnification. Head view of the front of a harvestman. Photo A is focused on the eye and only part of that is sharp, because depth of field can be measured in microns. Very little else can be seen, let alone is in focus. Moving to refocus further forward, on the tips of the mouth palps, photo B shows these in focus but like before, everything else has disappeared. This makes setting up a composition before focus stacking difficult. (Raynox 505 lens mounted on Canon 70–200mm lens at ƒ22.)

Vibration

As well as the magnification of the subject, there will be an enlargement of everything else, from dust and particles of debris on the eye of your insect to problems with subject itself. Most important will be the increased chance of vibration and movement. Outdoors, the slightest breeze will move insect wings and stamens, not to mention a leaf with your subject sitting on it. Everything needs to be anchored down and suggestions for this will be given later. Even if you can control the subject, one of the worst problems is vibration of all the equipment. Identifying why an image is soft can be difficult to pin down. Using live view on the camera or preferably when it is relayed to a larger screen helps, especially if the latter enables you to control the system remotely to reduce vibration. However, when you have problems, analyze everything from shutter and DSLR mirror movement to traffic passing by outside the building. Working on a solid table or work surface is essential but if that is located on floorboards, expect vibration if someone is walking nearby.

Sensor Dust You may think your camera sensor is fairly clean, as no discernible dust spots appear when you use your normal photographic lenses. Even a standard macro lens seems fine. Once you are magnifying subjects beyond macro, black spots may become clearly defined. Try to minimize the removal of lenses as much as possible, always switch off the camera before doing so and perform lens exchange in as dust-free an environment as possible. Ultimately be prepared to clean the sensor fairly frequently and there may be the need to use the clone tool in photo-editing software later down the line.

Chapter 2

What Cameras are Best?

martphones can take remarkably good photographs and much of the hype when a new one is launched centres around the features of the camera. It has become all too clear in recent years that the cheap, compact, point-andshoot camera has all but disappeared. In turn, this has led to some major problems for manufacturers and even their demise as they failed to see the change in trend to people using camera phones. If smartphones are so good, why do we not see wedding photographers using them instead of complex digital single lens reflex (DSLR) cameras?

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Fig. 2.1 Pin mould on a tomato. A tomato had split in the fridge and quickly developed a furry coating. (×6 magnification, Canon 7D with 65mm MPE lens, ƒ8 and macro flash. Composite of forty-five images stacked.)

IS THERE AN IDEAL CAMERA FOR EXTREME CLOSE-UPS?

Much revolves around a camera’s lens size and the problem of vibration. As we will see, there are a number of methods to achieve close-ups and if you require a wide range of magnification for a diversity of subjects, you may find it difficult to find just one camera to do the job. Camera sensor resolution, the number of pixels in a photograph, is not a problem with modern digital cameras as they now have more than enough. High resolutions, such as 50 megapixel, require high-quality lenses as all those extra pixels will show up any imperfections in the glass. Additionally, for many of the lenses used in microscopes, the glass will not resolve high pixel counts and so they become largely irrelevant. We have seen that APS-C sensors can be perceived as having some advantage as the image result has a higher magnification, although a full frame sensor will have a higher quality image that could be cropped. In most cases, a camera that allows the removal of the lens is most useful. This means both DSLRs and Compact System Cameras (CSCs) are appropriate. But what about the smartphone? USING SMARTPHONES Costa Rica is a great country for photographing wildlife and is a favourite of mine. It is typical to go out into the rainforest as the sun is rising with a local guide who can find and identify some of the spectacular bird species. Armed with a telescope on a tripod, they can quickly set it up and point out a species at the top of the tree canopy no one else could see. It was some years ago, on my first trip, that I saw guides hold their mobile phone lens against the eyepiece and take photographs. Holes on the forest floor can be home to tarantula spiders. These and other invertebrates were also tackled with the mobile by putting it against the lens of reversed binoculars. Ten years on this has become quite mainstream and a range of products can be purchased to attach a smartphone for close-up photography or clamps to hold the device to telescopes. These are not just cheap plastic structures; some gadgets produced by companies like Sony and DXO are high-end products matched with quality support. The reason this is all possible is because of the small lens on the smartphone, enabling it to cover the small lens found on telescopes or microscopes.

Smartphones Despite the small sensor they can produce amazing quality images and should not be dismissed. The small lens has a real advantage in that it can be held over the eyepiece of microscopes and other magnifying devices. So useful is this that a variety of attachments are now available specifically for this purpose. Additionally, there are clip-on microscopes or magnifiers available online. Those advertising magnifications up to ×200 may be pushing the quality somewhat but typically for the cost of a round of coffee, there are some able to produce good record shots at ×80.

Fig. 2.2 Phone microscope. The clip holds the eyepiece over the lens of the smartphone. The integral light is switched on and the subject held close to the end. Focus and zoom controls are on the side. Fig. 2.2 illustrates one very simple device that is small to carry around. It is of metal construction with glass lenses and magnifies between ×60–100 using a zoom facility. In their own right they are good magnifiers and before connecting to a phone, test it without the phone. To do this, remove the clip that holds it to the phone (usually they unscrew). Buy one with a built-in LED light. The specimen needs to be brought very close to the base of the microscope and the clear plastic guards will help to hold the material roughly in place. Switch on the light and rotate the zoom control so that you work initially with the least magnification. Once you are used to operating the

microscope, reattach the clip and place the eyepiece of the microscope over the camera lens. You may need to move this around a little while watching the phone screen to ensure a complete circle appears. If the camera lens is not exactly over the eyepiece, the image will be distorted. About now is when you need at least three hands. A support to hold the specimen helps while one hand supports and moves the phone carefully to focus. The other hand takes the photograph. Resting elbows on a table is almost essential, as magnification of even the lowest ×60 will exaggerate movement. Not only is it tricky to hold still but like most microscopes, optically, everything is in reverse. The light may be coming from the left but when you look through it, it appears to be entering from the right. This can be confusing until you become used to it.

Fig. 2.3 Anther of a rosemary flower taken with the phone microscope. (Estimated magnification ×60.) In practice, to achieve a reasonable image, take many photos and perhaps one in ten could be acceptable. The act of touching the screen to fire the shutter is enough to move everything. Try putting the camera in timer mode or if there is a burst mode available, hold your finger down on the screen and take multiple shots, trying to hold everything in focus. This technique will work with compound microscopes, as discussed in Chapter 4. In the case of all microscopes, the camera lens is wider than

the eyepiece and so vignetting occurs, creating a circular image with black in the corners. Normally digital zooms are frowned upon but in this scenario, zooming into the picture on the screen will enlarge the circle so that it touches the screen edge and reduces vignetting. On the computer, the image can be cropped and enhanced. Smartphones can therefore have their place in the photographer’s kit. Not only is the camera component useful but the phone can also be linked with a more sophisticated camera by Wi-Fi or NFS. The image can then be made to appear live on the screen to aid focusing, control the camera remotely and trigger the photos to be taken without touching the camera. This reduces vibration.

DSLR cameras These cameras are the type that we might assume are the best. Most importantly the lens can be removed and others more appropriate for the work fitted. Not only do these cameras have a system of accessories supporting and backing them up like flash and macro lenses, but a huge array of third party material can be purchased that will work with them. This applies especially to Canon and Nikon. Pentax may not be quite so straightforward but with a little persistence achieves results just as good as the others. Sony has a large collection of camera bodies but these are not strictly in this category and they better fit the next section. A true DSLR has light passing through the single lens at the front on to a mirror, which reflects the light up through a prism on the top of the camera, exiting through the viewfinder at the back. This mechanism takes up space and is heavy but the view is an optical one, giving a faithful rendition of what the lens sees. In the Sony A series of bodies they are called SLT, as the mirror is semi-translucent where a third of the light passes through to the sensor for autofocus and the remainder for imaging. There is no optical viewfinder but an electronic version. What DSLRs do have is a live view so that with the flick of a switch, an electronic view appears on the rear LCD screen. They therefore have the best of both worlds. We will see that for extreme close-up work the live view is close to essential. DSLRs are very controllable and with the right software, all functions can be operated remotely, a feature not typical of other cameras. The potential weak point of the DSLR is, however, that mirror. As the shutter is fired, so the mirror has to lift to allow light to pass through to the sensor. As the mirror rises and stops, vibration will occur through the camera and therefore the sensor. At the end of the exposure the mirror returns, again causing this ‘slap’ action. It is the first slap that is worst for any slow exposure and will be magnified by the extreme close-up methods. The higher the magnification, the worse this problem can become. We will see that using a flash can remove the problem but fine detail can be missing due to the slap. All DSLRs have a mirror lock-up facility available through the menu to help negate this problem. As the name suggests, the mirror is locked in place to prevent the slap. Except there is another problem: the shutter itself. The

focal plane shutter mechanism employed consists of two curtains. The first is across the sensor and when the exposure is made, it is withdrawn so that light reaches the sensor. To stop the exposure, a second curtain comes across to block the sensor. The first curtain movement can cause a possible vibration of the sensor at slow shutter speeds around a quarter of a second. Long exposures beyond several seconds or of shorter duration have no obvious sign of vibration. This is mainly visible on high magnifications but should be considered when assessing photographs.

Fig. 2.4 Reproductive structure of the lichen, Cladonia. Note the image is very soft due to first curtain shutter vibration. Composite of 32-image stack using LED lights. (×7 magnification, Canon 7D mk2 with bellows and objective lens.)

Fig. 2.5 As in Fig. 2.4 but by taking the photographs in live-view, the first curtain shutter is replaced with an electronic one that does not produce the vibration, resulting in a much sharper image. Professional and enthusiast Canon cameras have several advantages here, although Nikon cameras are beginning to appear with similar features. Many Canon cameras have a ‘soft’ feature, where the mirror has special dampers that soften the slap action. Additionally, in live view, the first curtain mechanism is replaced by an electronic shutter; effectively a switch that activates the sensor. When live view is engaged, you hear the first shutter open, waiting for the electronic system to come into play. SHUTTER ACTUATIONS One possible aspect worth considering when choosing a camera for extreme close-up photography is the reliability of the shutter mechanism after a large number of actuations. In general use this is not necessarily an issue. As we will see, focus stacking involves taking a huge number of photos in a relatively short time; I can easily fire the shutter a thousand times in an afternoon. In the product information, a manufacturer will typically say how many actuations the

camera shutter can take before it may become liable to problems. Relatively expensive DSLR models will be listed at around 200,000 actuations.

COMPACT SYSTEM CAMERAS (CSC) OR MIRRORLESS CAMERAS Panasonic brought out a new type of camera in 2008. Digital still photography has its origins in video and these manufacturers arrived a little late to still photography and needed to develop their own niche area. The design of DSLRs was already well advanced by others like Canon and Nikon. Sony, another manufacturer rooted deep in video technology, cut a quick advantage by buying out Minolta-Konica and so had cameras and lenses already well developed, ready to adapt. Competition for new cameras was so strong that just like in nature, as new designs appeared, some were to become extinct while others thrived. Olympus has always tried to produce small, lightweight cameras and joined Panasonic in the new Four Thirds sensor. Smaller than APS-C, it allows for small but high-quality cameras. Fuji had a good DSLR using the Nikon mount for lenses but suspended production and switched to a smaller design using an APS-C sensor. Together these companies produced a new market for quality, interchangeable-lens cameras. Nikon developed its own CSC based on a small 1in sensor and although it had a reasonable following, was effectively discontinued by the beginning of 2017 as the sensor cannot compete with rivals. Canon was the last to the CSC party with the petite M series. Using an APS-C sensor, the quality is excellent and up with the rest. These CSCs have several advantages over the DSLR. Dispensing with the heavy prism and shutter mechanisms, they are smaller and yet have similar sensors. The image quality is arguably as good and yet without the mirror, there is no slap with a minimal amount of vibration. One dispute over their viability was the delay between seeing the subject in the electronic viewfinder, squeezing the shutter release and the photo being taken. Recent models show this to be a negligible issue with electronic viewfinders having controls to speed up their refresh rates so that the screen can keep up with fast movement. Being lens-interchangeable, they have the ability to attach any lens either for their specific mount or any other using an adapter.

Fig. 2.6 Peak focusing on a Sony A6500. The red areas identify the pixels in focus, a helpful guide especially in this case on a microscope. Sony produce two series of mirrorless cameras. The A mount series is for fullframe sensors and the bodies are larger than most CSCs. With the high-quality sensor, transparent mirror and electronic shutter, they have become a firm favourite for photomicrography. The alternative E mount series use APS-C sensors, have no mirror at all and also have the possibility of an electronic first shutter mechanism to minimize vibration. A clever feature found here and other CSCs, for example the Panasonic cameras, is peak focusing. There is a choice of low, medium and high sensitivity and you can alter the colour. On the camera’s rear screen (not the viewfinder) any sharp, in-focus patches or lines of pixels show up as red by default. For general use, like landscapes, this can be distracting but for extreme close-up photography where achieving focus can be so difficult, it is an incredibly useful feature. When focus stacking using the camera’s live-view, peak focusing enables a much smoother and quicker set of stacks. Instead of moving the camera trying to check for focus in the subject each time, one just needs to monitor the changing red;

move the camera watching the red shift position slightly on the screen and then take another photograph before moving on to the next. Connectivity to computers, smartphones and tablets has been a problem for CSCs in the past. All have a Wi-Fi connection to a phone or tablet so that an app permits some control of the camera. This can be somewhat hit and miss, varying between makes. The usefulness to us is to have live-view on a larger screen and control the camera to take photos that can be instantly downloaded. Full control is typically lacking and is the one area in which DSLRs excel. A larger screen for working with extreme close-ups is very useful and there is more on this at the end of the chapter.

LENS ADAPTERS A quick search on an auction site like eBay will list plenty of adapters that allow any lens to be fitted to any camera body. This has been a growing trend for the last five years and has gone from a very niche position, only available from obscure Chinese sellers, to the large lens manufacturers. This has driven prices down considerably although there is a huge variation from a few pounds or dollars to many hundreds. So what are the differences? The lens sends information through electrical contacts to the camera body so that focusing and apertures can be controlled. The more expensive adapters will have these contacts (automatic) but if attaching a camera to manual bellows where no electrical connection is necessary then a cheap, manual adapter is all that is required. Adding an adapter tube between the lens and body could extend the lens so that infinity focusing is impossible. Again this is not a problem for close-ups but if you intend to use the adapter with a lens to reach infinity, buy a more expensive one.

Fig. 2.7 Camera adapters. The Minolta-mount bellows has a manual adapter attached for a Canon DSLR (MD-EOS). The Sony 6500 has an E-mount automatic adapter suitable for Canon lenses and therefore enables the Sony to fit the bellows. The best (and unfortunately the most expensive) of these are produced by Metabones and Sigma but have all the features necessary to enable the lens to talk to the camera. Significantly cheaper ones may do the job but the speed of focusing is impaired, becoming sluggish and often hunting back and forth. However, for macro and extreme close-ups, focusing should be done in manual and autofocus switched off. As we will see in the next chapter, some of the methods we use have lenses with no aperture control. For these, the very cheapest adapters can be bought where it is simply a ring with the correct mount on either side. Lens adapters have made our photographic lives much less complicated. No longer do we need to stay with a specific manufacturer but can buy lenses and cameras to suit. There is no excuse now to have a multiplicity of cameras in your arsenal!

Fig. 2.8 Olympus TG-5 compact camera with the LED light guide fitted.

IS THERE STILL ROOM FOR COMPACT CAMERAS? The basic, point-and-shoot compact has all but disappeared under a barrage of smartphones. In fact, Nikon announced the closure of their Chinese factory for this reason in November 2017. Innovation, however, has produced a diversity of small cameras for niche areas. One development of the small sensors compact has been the burgeoning numbers of waterproof, shockproof go-anywhere cameras. While this has been aimed at the activity sports person, they have very close focus, due to the small sensor size, and are brilliant to keep in the camera bag for occasions when the weather deteriorates, for underwater photos or a quick close-up.

Fig. 2.9 Images, all underwater in rock pools, taken with the TG-5. A: beadlet sea anemone; B: limpet with shell covered in green gut weed; C: acorn barnacles; D: snakeslock sea anemone. One model stands out from the rest: the Olympus Tough TG-5. Unlike other waterproof compacts, there is some control over exposure with the option of RAW format and aperture priority. By selecting ‘microscope’ mode, there is an option for the camera to focus very close to objects providing a magnification of up to ×4, although due to the small sensor size this is not beyond true macro. However, it is impressive. Light is an issue when very close, as subjects will be shaded by the camera body. A small LED light guide accessory can be fitted which directs light from an LED, located next to the flash. By setting this up on the menu, the light remains on and gives a reasonable breadth of light on the microscopic subject. Upon selecting the microscope mode, several other options appear including focus stacking and focus bracketing. The former is an automatic stacking sequence where the camera takes around fifteen to twenty shots in very quick succession followed by a short ‘busy’ message while the camera creates the composite image. This is saved

along with the first in the sequence. There is a minimum amount of control in the stack and it is best to make sure the lens focuses clearly on the main subject as not everything in the viewfinder will be picked up and photographed. By contrast, the second option of focus bracketing is a more controlled process. Upon pressing the shutter release, the camera takes a quick succession of images starting out of focus on the nearside of the view and as you watch the screen, the point of focus is followed until after about twenty-nine shots, the point of focus has left the view and the camera stops with all photos saved. You process the composite later using suitable software on a computer. Here, the entire field of view is stacked. Although you will need to keep as still as possible while the camera carries out these operations, the process is very forgiving and allows for a surprising amount of leeway and camera shake. Both methods take nine to ten seconds to complete. Of course the real ace is that this can be done underwater. Just by holding the camera in a tide pool on the seashore or in a pond, quite amazing photographs can be taken. The battery on many compacts can be quite poor but the TG-5 is remarkably good with an afternoon shooting of thirty to forty sets of focus bracketing (1,160 images recorded) easily possible. The previous model, the TG-4, had very similar stacking options but was slightly slower. Olympus has focus bracketing in a number of the Micro Four Thirds CSC cameras but of course they lack the flexibility of the TG-5 in doing this underwater.

BRIDGE CAMERAS Before the DSLR became more affordable, the bridge camera developed a very good name, bridging the gap between compacts and the more serious DSLR. With the advent of the Canon 300D in 2005, the first ‘cheap’ DSLR, fewer manufacturers put finance into their development. With similar sensors to the compact they became popular due to the good zoom lens and controllability. In recent years there has been a serious resurgence, especially by Sony, with better sensors and excellent lens performance. Although the lens cannot be removed, they can fit the category of cameras that can have lens attachments added in front of the front element, in particular using the technique called ‘coupling’ seen in the next chapter. The zoom lens is useful with this method to increase the magnification and the larger sensors provide good quality images. The term ‘block’ has been used more as a descriptive nickname. Fairly new to the scene, they lack a general category name. As this name suggests they are a block, often a flattened cube containing a sensor. Principally, they are designed as a camera for certain microscopes fitted to the top of the microscope tube. Some have controls on the top of the block for switching on/off, adjusting exposure and taking

photographs or video. Others have no controls and so need to link with a computer where dedicated software controls the camera. The latter is the most typical arrangement for fitting to microscopes and pricing varies considerably. A live-view within the software allows adjustments to be made and photos taken that are stored on the computer. The cable from the camera is connected to a USB port and obtains its power from the computer. The former type of camera that has controls is a relatively new and inexpensive development. When searching for them on the internet, note that they are often referred to as ‘industrial inspection’ microscope cameras. There is quite a diverse range of possibilities and most will come with extra kit which varies from just leads up to lenses and stands, lights and even remote controls. The latter is a good option as taking the photos remotely prevents shake. These are discussed more under the section on USB microscopes.

IMPORTANCE OF LIVE-VIEW AND TETHERING With the problem of focusing in extreme close-up photography, looking through a small viewfinder becomes increasingly tedious and difficult. Just a gentle knock with your head on the camera can put everything out of focus and you will need a remote control for the shutter so it will be hands-free. Sitting back from the camera and using live-view is preferable. The screen on the back of the camera is better than nothing, but is typically too small. Most cameras allow you to zoom into the live-view screen, enlarging areas to check for focus. Touching any camera controls will cause vibration and as well as live-view, consider ways to operate the camera remotely, both for working indoors or out in the wild.

Fig. 2.10 Wireless live-view in the field on a tablet. Here, the tablet is connected by Wi-Fi to a Canon M3 so that the focus stacking operation on a lichen can be completed without touching the camera and causing vibration. Virtually all the cameras mentioned above have phone and tablet apps that link wirelessly via Wi-Fi, Bluetooth or NFC. Some are noticeably better and more stable than others so if this is important to you, it may be worth checking the availability and reviews of remote apps before you buy the camera. From the tablet or phone you should be able to see the live-view, change at least some features such as ISO and

aperture and take the photograph. Most have a choice for the image to download on to the device that may be edited and sent to social media if you wish. Focus stacking is not really an option on phones and tablets although Helicon Remote is a third party app that controls Canon and Nikon DSLRs remotely but stops short of actually stacking images. USB connections are reserved for laptops and desktop computers. Live-view can then be relayed to large screens, for example a 32in 4K monitor that provides unrivalled ability to find focus. A USB tethered camera will have access to all the software applications you might need as well as the live-view. Now the problem. While all cameras come with the leads and ability to download images to the computer, the software to see a live-view image and to fully control the camera remotely may not. Canon cameras are packaged or have support links to freely download the necessary software, called EOS Utility, that is specific to the camera model. Nikon users may have to purchase the software for full control. However, Helicon Remote is a good third party option. If you use the photo editor Adobe Lightroom, there is the ability to tether a camera with limited control and live-view. Other camera users can struggle, particularly CSC/Mirrorless ones. One possible option to consider is Capture One, a photo editor to rival Lightroom, although you do need the full version, not the one listed as Express. A trial version can be downloaded and tested before you need to pay but can be well worth the cost. Live-view can have a downside in that it eats up the battery power. A DSLR will struggle to last several hours on a full battery charge when performing a good deal of focus stacking. That may seem fine but it is surprising how long it can take and a battery warning in the middle of a stack is the last thing consider buying a mains adapter for the camera. If you regularly want to spend a day or even a morning stacking, they are highly recommended. Some CSC cameras, for example Sony, that charge batteries while they are in the camera, can use the charging adapter to power the camera.

Fig. 2.11 Underwater close-up in a shallow pond with the tiny carnivorous plant bladderwort, all under the leaves of bogbean. (Olympus TG-4 held under the water and the shutter released.) In addition to a USB connection, an HDMI link might help. This is a good alternative for some CSCs lacking the USB control facility. The HDMI can link to a small screen monitor and will be a useful help in the field when sharp focus can be elusive. These 7 or 9in screens run between 9–12 volts and can be attached to a mount connected to a tripod. Smaller and larger monitors are available but these sizes are the most practical. By checking around online, you could find deals as low as £40 — or up to over £1,000 – that get you the state of the art with additional functions. Either a tablet/laptop screen or a small monitor will make focus stacking much easier to do. The latter is small enough to be less intrusive or as potentially problematic as taking a tablet into the field but only provides a live-view, no control. Ultimately, it is down to personal preference and what works best for your circumstance. WINDOWS TABLETS Most tablets and phones commonly available are for Android or iOS operating systems. I have

used an iPad wirelessly with a Canon app and an M3 CSC, effectively creating extreme close-up stacks in the field. However, I often would like to check the composites at the time and have full control over the process. Tablets for Windows are not so abundant and can be expensive, like the Microsoft Surface Pro series, or value ones that are the most inexpensive tablets to buy. I have used both when working in the field and they are excellent. The most important feature for me is that by having a Windows OS, the full camera software like EOS Utility is there to operate my EOS DSLR, not a basic app that may or may not connect wirelessly. I use it connected by USB 3 cable and can stack the images if I need to. In other words it is just like a tethered stacking process at home. My concern over the superb Surface is the expense if it gets wet or dropped, although the screen is much better in the field. Expensive cases can be purchased but for the last year or so, I have had a cheap Tesco Windows 10 tablet and a cover I bought on eBay all for less than £90. Memory is limited and so a fast micro-SD slot has a card receiving all the stack images from the camera as they are shot. If composites are made, these go on there too.

Chapter 3

Techniques to go Beyond Macro

alk into any camera shop or check out the merchandise online and straight away you can buy a macro lens to produce life-size images of wildlife. Focusing from infinity down to 1:1, the lens will potentially enable you to achieve good close-ups in no time. Going beyond macro is not so straightforward. You have entered an exciting realm of specialism but it need not be expensive. Methods are diverse and constantly evolving. Just be prepared for a great deal of trial and error but hopefully, after hours of playing around, a eureka moment will make it all worthwhile. At the end of the chapter is a table to summarize the majority of these techniques.

W

Fig. 3.1 A marine worm found living in the base of a seaweed. Marine worms are very different to those on land with a huge biodiversity. This one is just a few millimetres long but some species are up to a third of a metre in length. This was a live specimen on a microscope slide, in seawater with a coverslip over the top, photographed with one flash on top and one below. (×7 magnification, Canon 7D mk2, 65mm MPE lens and extension tubes, ƒ5.6. Stack of five images with some retouching as the worm moved.)

CHALLENGES Some of the challenges around extreme close-up have already been discussed, such as vibration, light and focusing. Remember from the start to switch off autofocus as everything will be in manual from now on and focusing will be done by moving the camera (or subject). Reducing vibration by having a stable base and light is so crucial to success that they have their own chapters. Two practical issues before starting the techniques: working distance and flat field. The first of these refers to the distance between the end of the lens and the subject. As magnification increases, so the working distance decreases. The problem that creates is the ability to get light on to the subject. How limited this distance is will depend on the lenses and methods used, so requires consideration. Flat field is rather more complex to understand. The whole point of using lenses in photography is that they alter the way that light travels. Passing in straight lines into a lens, the curvature of the glass changes this pathway. Called refraction (reflection bounces it off the glass surface), the course of the light is now bent. Convex lenses (fat in the middle, thin around the edge) bend the light to a spot, the focal point, while a concave lens (thin in the middle, fat around the edge) bends the light outwards. A convex lens is the one that provides magnification but the key problem is that the middle focuses at a different point to the edge. This means that while the middle gives you a magnified image, the edge is out of focus. This is spherical aberration, a distortion due to the curvature of lenses. Now we see why a camera lens has a number of elements, not just one convex lens but a mix of these and concave lenses to correct the image and hopefully provide an even and sharp photo. Although more expensive to produce, manufacturers use some critical elements in their lenses to be made from fluorite, producing clearer images. Magnification can increase these problems and such lenses are corrected for close focusing and this is where we look out for lenses that have a flat field and minimal spherical aberration. Another aberration that we see associated with the techniques here involves colour.

Fig. 3.2 Parallel rays of light are brought to a focal point behind a convex lens but they focus at different points, causing spherical aberration. White light passing through lenses (or any transparent object from water to plastic) can be broken up (diffracted) into its component wavelengths, particularly the primary red, green and blue. All lenses are potentially prone to it but with high magnification, the last thing you want is a rainbow around the image. Typically, this chromatic aberration manifests itself as fringing to strong edges in the subject. Computer software may remove a very mild form of colour aberration and fringing but it would be better to reduce it in the photograph in the first place. Selecting lenses for extreme close-up can be a minefield and often you do not realize until using a lens that you have a problem. An impractical guide to this is the more expensive the lens, the more corrected the lens is likely to be. A more helpful indicator is looking to see if the lens is described as achromatic, apochromatic or planachromatic. This should be written on the objective lenses used in microscopy and may be provided in the data for other glass magnifiers. In essence, the achromatic lens is corrected for red and blue while apochromats have green corrected as well. The latter can have fluorite glass (higher quality) elements and these are referred to as fluorites or semi-apochromats. Plan

versions are the most expensive lenses with colour correction and additionally, these correct for curvature or spherical aberration by producing a flat field. When buying any new photographic lens, it is worth testing to see which apertures produce the sharpest picture and where any distortion or aberration lies. A lens test sheet (downloadable versions to print are available online) is hung up in good light and with the camera on a tripod, this is photographed at each full aperture with results analysis enlarged on a computer screen. An alternative to a test sheet is a white A4 sheet printed with narrow black lines. Check that the lines are sharp and straight but especially that when enlarged there is no colour distortion. Lens correction profiles can be downloaded or created and saved in Lightroom.

EXTENDING THE LENS Take the lens off your camera body and look at the rear element. This is often referred to as the projection lens as it is the glass to project the image on to the sensor. The latter is rectangular and makes a rectangular photo from a circle of light passing down the lens. Depending on the chosen aperture and lens, the corners of this rectangle are so close to the edge of the circle they may darken slightly. This is called vignetting and is a common problem. The light being projected from the rear element is actually a cone. To view this, there needs to be a bright light entering at the front of the lens; dim the ambient light and an image can be seen projected upside down if a piece of white paper is placed behind the lens. Move the paper slowly away from the rear of the lens and you will see the image enlarge, but as it does so it will lose brightness. Let us presume that the lens is a macro set to 1:1, life-size magnification. Within the camera, the sensor acts as the screen to collect the image by grabbing the photons of light as a rectangle shape in the middle of the cone. The image is 1:1. If the camera body is detached from the lens and moved away from it, the projected cone has a much wider diameter and if the sensor is exposed, it takes a small proportion of the projected image, which on the sensor looks much larger. The further the distance between the rear lens element and the sensor, the greater the magnification. If you start with a 1:1 (×1) image then the magnification, in theory, is unlimited but the amount of light diminishes with distance. Secondly, the image quality will eventually deteriorate as any faults in the lens will magnify as well.

Fig. 3.3 An extreme macro set-up using a Sony CSC with extension tubes, lens adapters and a microscope objective lens mounted on a stable base to photograph fungal threads on a twig at a magnification of approximately ×14. Two homemade LED spots illuminate the subject. Note the limited working distance: the distance between the end of the lens and the subject.

Fig. 3.4 Extension tubes. Third-party tubes are considerably less expensive than those from the camera manufacturer, which in the case of CSCs are not made at all. If you already possess a macro lens then the first method to increase magnification will be to extend the lens. It is not sufficient just to take off the lens, as the image degrades with stray light entering the camera. A tube made from drainpipe tubing and painted black inside will work, with one end fixed to the camera body and the lens fitted at the other end. Plus plenty of light on the subject. It does work but is very difficult to control.

Fig. 3.5 Crab spider. Using a CSC with two sets of extension tubes and a flash, the spider was photographed from the front, using 35-focus stacked images to create a composite. (×4 using a Sony A6500.)

Fig. 3.6 Bellows. These are a generic set branded as Unitor from the 1980s with a Minolta mount. Here, a manual adapter enables a Canon DSLR to be fitted and at the end, a Vickers objective lens. Extension tubes are available in an incredible variety of prices and qualities. As it is just a tube, there is no point buying the manufacturer’s own version. Commonly available in camera stores are third-party brands. Even cheaper ones are produced in China and on sale through most online retailers and auction sites. Made from plastic, the inexpensive tubes sell for between £9 and £20. These are automatic, meaning they link the aperture diaphragm to the camera. If you are using lenses without apertures then manual ones at even cheaper prices are available. At this low cost it is worth having more than one set of tubes. They do vary in quality and the ones with aluminium ends to each ring give the best rigidity and connection. Some good and inexpensive names include Neewer, Meike and Pixco. Extension tubes can be bought for any interchangeable lens camera system, but for CSCs only these inexpensive third-party options are available. Check the tubes out with a variety of lenses that you may have. A telephoto usually works better than a wide angle but just check and see. Tubes work with most of the lens options we will discuss further on in this chapter.

Fig. 3.7 Head louse (dead) photographed with a Canon M3 fitted with a 200mm lens and a 50mm lens reversed and coupled to the front. (×4 magnification, sixteen-image focus stacked to create a composite.) Bellows are another way of extending the lens. They are not so widely available as extension tubes but provide up to 200mm of extension while a set of three DSLR tubes of three sizes will be 13mm, 21mm and 31mm respectively. Although automatic bellows (to change aperture) can be found, they are not worth it as invariably they will be used with a lens having no diaphragm. Buying second hand bellows is fine but check they have no holes as these will let in light and even the tiniest amount will fog the image, reducing contrast. Bellows work well with camera adapters and so a set with a Canon mount can be made to fit any other camera with the right adapter.

DSLR LENSES, REVERSING LENSES AND OTHER CAMERAS Even if you have no specialist lenses there may be something you have that will allow you to go beyond macro. By taking a lens off the camera and reversing it over the

body, an image beyond macro is possible where the quality glass is just a super magnifier. Adapter rings are very reasonably priced, as there are no connections to be made other than the screw filter thread and camera mount on the other side. Wide-angle lenses provide the greatest magnification. The diaphragm needs to be used wide open to receive enough light to see. This is not controllable anyway and can be a problem if the lens defaults to the smallest aperture on removal from the body. An arguably better way of using a reversed lens is through a system of coupling whereby the reversed lens is mounted on to the front of another lens, preferably a telephoto. This is an ideal method for bridge cameras where the lens cannot be removed. A standard 50mm lens from an SLR camera can be reversed and attached via a coupling ring. If the filter threads are not the same, you will need one or more step up/step down rings so that it fits. Always try out a lens combination first by just holding the reversed lens in place with your hand while you move the camera back and forth to focus. Note that many people when first trying this technique (and others like the Raynox lens later) say they cannot see anything. This is because they have not got close enough to the subject. The working distance can suddenly drop to just a few centimetres. This coupling technique works well with any camera, even compact cameras, and is an excellent way of starting out with extreme macro. If the 50mm lens is reversed on to a 300mm lens, the magnification is more than ×6 (300/50=6). This is the situation with the CSC camera in Fig. 2.10, photographing a lichen on a rock in the field.

Fig. 3.8 Raynox 505 Ultra Macro Converter fitted to a 70–200mm telephoto lens.

ATTACHING EXTRA GLASS No matter how glass is made there will be some form of distortion, whether these are changes to the image shape, focus is skewed or colour altered in some way. Manufacturers spend a great deal of time trying to minimize this when producing camera lenses. By adding more glass elements, we can increase the magnification of a lens but expect the quality is likely to be reduced in some way as we alter the passage of light. For many decades, screwing a close-up lens into the filter thread of a standard lens on the camera has been an option. Coming in 1, 2, 3 and 4 diopters (a measurement of their strength), these are like holding a magnifying glass in front of the camera. Magnification is not exceptional and for the aim of this book, not particularly useful. One worth mentioning is the Canon 500D close-up lens – but it is expensive. They are easy to carry in a pocket and are popular with birders with long focal length lenses. Designed for a 100–400mm lens, they bring the minimum focus down to virtually macro.

Cheap, single elements of glass can particularly lead to chromatic aberration and edge-to-edge sharpness will be missing. Stopping down to ƒ8 or ƒ11 will help as less light is taken from the edge. To improve on aberration, some add-on glasses come as a compound attachment constructed from several elements of glass. These are generally called a macro converter lens, which also attaches to the front of the main camera lens. The Opteka 10× is one example, listed as an achromat lens, which receives reasonable reviews and is a low cost option, screwing into the filter thread.

Fig. 3.9 Head of marine worm which is emerging from its burrow taken with the Raynox 505 on a 200mm zoom lens set to ƒ22, flash from above. (×12 magnification with live animal in small observation cell.) Perhaps the most widely known of these converters is the range of Raynox macro conversion lenses. This Japanese company has been making lens attachments for

over thirty years for both still and video cameras, to include extreme wide angle or telephoto. Steadily increasing the macro range to four, they start with the 150 and 250 converters that give true macro performance. Then there are the pair of Ultra Macros, capable of going beyond macro and because the attached converter fits to the end of the lens, they can potentially work with any camera. The 202 model reaches magnifications of around ×6 while the 505 is their most powerful and can be pushed to ×15. Achieving that kind of magnification is not without its difficulties. The Raynox Ultra Macro lenses have a screw thread of 37mm but it is easier to attach via the included adapter. Like the others in the range, the adapter is spring loaded like a lens cap and firmly clips to the filter thread of any camera lens with diameters between 52–67mm. It is recommended to use a telephoto lens and a 70– 200mm zoom is ideal. In use the greatest magnification comes with the longest focal length and provides a good working distance (2cm for the 505). To achieve the best result, the zoom lens requires stopping down to ƒ16 or ƒ22. This will create a problem of lack of light, which a flash will cure to give the sharpest images. Focusing can be really difficult and ultimately the only usable photographs will be those that are focus stacked on a focusing rail. In conclusion, the Raynox macro conversion lenses are good methods for magnification but the high magnification of the 505 makes it more difficult to use and produce really sharp images.

Fig. 3.10 Lens extender. The 150mm Sigma macro is fitted with a Sigma extender designed for the lens. In addition there is a full set of extension tubes. This produces ×4 magnification. The camera’s flash is raised to wirelessly trigger the flash (complete with homemade diffuser) held above the fly, indicated by an arrow on the wall (see Fig. 3.11).

Fig. 3.11 Predatory fly. This is the one being photographed in Fig. 3.10. Note the green dots on the wall; these are single cells of algae. (×4 magnification, ƒ14 with diffuse flash.) A very different way of adding glass is putting some between the lens and the camera body. These are called converters or extenders and are marketed as a way of increasing the focal length. However, a ×2 extender fitted to a macro lens on 1:1 magnification will be extended to ×2 and for APS-C even higher. The quality is likely to drop with the additional glass but a ‘matched’ extender, one made by the lens manufacturer specifically for the lens in question, will be the best. Also consider a

×1.4 extender instead as any imperfections in the lens will not be so enlarged. Sigma produce some very sharp macros in their EX range. The ×1.4 extender produces a super sharp image with, say, the 150mm macro, and if extension tubes are used as well, magnifications in the region of ×4 are of more than acceptable quality.

SPECIALIST MACRO/ MICRO LENSES Going beyond macro, there is really only one stand out lens, and one that is discussed more than any other online: the Canon 65mm MPE lens. Although classified as a macro lens, it actually starts magnifying at life-size. It is a non-focusing lens as this is achieved through moving the camera or subject. The only adjustment is to increase magnification up to 5:1 on a full frame sensor, 7:1 for an APS-C. With an aperture range of 2.8 to 16 you might be tempted, especially at the higher magnifications, to use ƒ11 or 16 but the results are poor. Diffraction – distortion caused by the way light passes through a small hole – increases to an unacceptable level. F5.6 is a good working aperture. A set of extension tubes will take the magnification up to around 12:1 before it becomes unmanageable. No other easily available lens is like it unless you consider the Zeiss Tessovar. Built in the 1960s as a microscope for low magnification, they appear on eBay very occasionally and have a similar cost to the Canon MPE. The Tessovar has good flexibility, with a rotating turret and several objective lenses for different magnifications. Working distances are excellent. Any camera body can be fitted with a suitable T2 mount adapter. Likewise any camera body can be fitted to the Canon MPE with the correct adapter. The Tessovar was only produced by Zeiss for a year or so before they ceased production as it was so expensive to make. At the time you could have bought a new car for the recommended retail price. The Canon optic does not appear to be in short supply, but many in the extreme close-up community believe that they also stopped production due to cost. Initial costs of the lens were so high when it was launched ten years ago that few were sold. After a number of years the price dropped to its current level. So there may be a finite number out there. Time to buy a second one!

Fig. 3.12 Zeiss Tessovar lens attached to the very heavy and stable Zeiss base. Despite the huge global CSC market the presence of macro lenses has been quite limited. With the large number of adapters available, existing optics for DSLRs can be used although a few of the camera companies are producing macros for their CSCs. They tend to be quite expensive although the Canon 28mm image-stabilized macro for their M series is not. Innovatively, it comes with a built-in LED ring light although the working distance is quite small. The Chinese optics company Laowa have launched a 2:1 macro that has reviewed

well. In fact there has been a recent boom in small Chinese optical companies due to their increased skill-set in lens making. High costs of production in Japan have pushed many camera companies to carry out manufacturing in China and so have passed on the specialist skill in optics. Looking for niche areas, several new Chinese companies have been developing special macro/micro lenses for mirrorless cameras. Zhongyi produce a super macro called the Mitakon 20mm ƒ2, which magnifies around 5:1 and with extension tubes will go to around 7:1. It is made for the Sony E mount and Canon M series. As it is fully manual, a basic adapter should allow it to be fitted to any CSC. At around £150 it is significantly cheaper than most options. Primarily purchased online to arrive from Hong Kong, there are now a few private sellers in the UK.

Fig. 3.13 Canon 65mm MPE lens fully extended, fitted to a set of extension tubes. All on an electronic focusing rail.

Fig. 3.14 Bellows fitted with a reversed Leitz enlarger lens. Yasuhara, whilst a fairly new Japanese company, produces lenses in China for the CSC market including the Micro Four Thirds sensor, Sony E and Canon M mount. One lens is the Nanoha macro ×4 to ×5 magnification which has auto apertures. It could be judged as the CSC equivalent of the Canon 65mm MPE and although not long on the market, is already gaining a European following as it is a sharp optic. What makes it stand out, other than it is half the price of an MPE lens, is that there is an LED lighting module that fits on the front of the lens and is connected via micro USB cable to a power unit. With working distance down to around a centimetre, this helps to direct light exactly where it is required. The lighting is more than adequate but has a limited flexibility and like all magnifications of this order, will require a great deal of stability. There is an EU distributor and it is sold at around 400 euros. With the burgeoning interest in macro and extreme close-up, new and innovative lenses are appearing. Information and reviews can be difficult to come by but worth watching out for.

LENSES FOR BELLOWS AND EXTENSION TUBES Enlarger Lens We have already considered using macro lenses on extension tubes. In fact, most lenses will take you to macro and beyond – but not that far beyond. The Canon MPE lens is the exception. Even the Raynox mounted on a 200mm lens can take a few extension tubes to push it just that little bit further. Thirty to forty years ago, when bellows were the only real option for even reaching macro as well as higher magnifications, manufacturers like Minolta and Olympus produced expensive, nonfocusing bellows lenses with a minimum number of elements and apertures for quality. The latter were fairly useless as apertures become redundant as you increase magnification creating tremendous diffraction (light scatter) and so sharp images were impossible unless the aperture was wide open. Bellows lenses are now only available second hand on auction sites, as are enlarging lenses. There has been a small resurgence in the popularity of developing and printing of film. Prior to this, second hand enlargers for printing the negatives were more reasonable in price than now, as we are seeing a slight price rise with the increase in home processing. An enlarger projects a flat object, the negative, on to a base to print on flat paper. All this ‘flatness’ could be a problem if spherical aberration was present in the lens. This is why enlarger lenses are flat field and will give very little distortion. Enlarger lenses have now become popular for extreme close-ups but there are clearly some better than others; for example, six element ones are better than four element lenses. They have apertures and the wider apertures are less distorted, so ƒ2.8 is better than ƒ4. All apertures are manual and the ring often has a gap in the lens where light passes through. In a dark room, this is useful so the aperture is highlighted. When fitted to extension tubes or bellows, if it has a hole, mask it with some black tape or fill with Blu Tack. Attaching to bellows requires an adapter or two, which are available for the typical thread size and mount. There has always been some debate as to whether the enlarger lens should be attached reversed or not as it is flat field. The best way to find out for a specific lens is to test it both ways. Also check the apertures. The likelihood is that it is better wide open. There is a reasonable diversity of second hand enlarger lenses on the market. The Leitz and Rodenstock ones are well known and good. Nikon produced a classic that has a cult following but make sure you buy the more expensive ƒ2.8 rather than the ƒ4 model. The Schneider Componon lenses are good, particularly the 35mm ƒ4 version. Objective Lenses This can range from a very cheap to a remarkably expensive operation; the variety of possibilities is high, which is good because ultimately this is the best method of

achieving high magnification, particularly when the likes of the Canon MPE finish.

Fig. 3.15 Selection of objective lenses. Note the black one at the back with an infinity symbol and on the right is the Minolta 12.5mm micro bellows lens.

Fig. 3.16 There are three microscope objective lenses here; a simple way to attach them to a bellows or extension tube will be to make a suitable hole in the centre of a camera body cap. We will see in Chapter 4 that microscopes have two sets of lenses, eyepiece at the top and objective lenses (often a number of them on a rotating turret) at the bottom of a tube or column. In this section we are just interested in the lower ones. Data about the lens is written on the side barrel and initially we need to know one of two things: is it a finite or infinite objective? If it is the latter, an infinity symbol should be printed there. The method we need to employ depends on this. Neither is necessarily better than the other, as this will depend on the quality of the lens. We have already seen that light passing through a single lens element will have various possible colour aberrations. Objectives have to be colour-corrected as on high magnification, the subject would be difficult to see with all the aberration. The degree of correction will increase cost but also the quality of the image produced. The basic achromat is corrected for red and blue wavelengths and spherical aberration is corrected in a green wavelength. An apochromat lens produces a higher

performance as it is corrected in red, blue and yellow while the spherical aberration is corrected for at least two wavelengths or more. In addition apochromats have a higher numerical aperture, which gives a shorter working distance. Numerical aperture, in essence, replaces the principle of the diaphragm aperture found on photographic lenses and is recorded on the objective barrel. Both these types of objective will suffer some distortion due to the curvature of the lenses, although it is more significant in high magnifications. This means that when the centre of the field of view is in focus, it degrades to the edge. A plan (or planar) achromatic objective can achieve more than a third better radial clarity from the centre than an ordinary achromat, as they are corrected to create this flat field. The magnification of the lens will also be printed, typically ×4, ×10, ×20, ×40 and ×100, although others do exist. In theory we could use any of these but for practical purposes (working distance being one) we will concentrate on the lower magnifications: ×10 provides a good and controllable magnification up to 40:1 attached to bellows. There are various makes to look out for, especially second hand, such as Vickers, Nikon, Olympus, Wild and Zeiss. Within these are many sharp optics at different costs. One of the most expensive sets of objectives comes from Mitutoyo, with a reputation as being among the best available, but occasional examples can be poor and care is needed when buying second hand. To help decide and search for good options, use a forum like photomacrography.net as it can be a real minefield. There are plenty of new, cheap Chinese objectives available but care should be exercised, as some are surprisingly good while others are appalling. This can apply to different examples of the same model, as quality control can be rather limited. To get around this, use a seller that has a good reputation, for example Joy Optics, and test immediately to check the quality. If there are problems you will then be able to exchange it. Second hand objectives from the well-known microscope brand names should be fine, but they need to be checked for fungal growths and a build-up of debris between the elements. Older objectives can have a problem of delamination where the element at the end over time has possibly come into contact with solvents from specimens. The edge of the glass can be affected but often does not change the workings of the objective.

Finite Objectives These objectives have a finite focusing point. This just means that as the light passes through the objective, the lenses bring the rays to a focused point on the other side from the subject (see Fig. 4.10 in Chapter 4). Attach them to the end of the extension tubes and bellows with the camera body at the other end and no other lenses need to be present. These days, mounting an objective is straightforward enough. Experimenting ten or so years ago was not so easy as adapters were not available. One method I developed, and still use on occasions, is to use a camera body cap. These come with cameras but are also very cheap on eBay. Find the mid-point and

drill a hole through the plastic. Check the objective but most are 20mm in diameter. By drilling a 19mm hole, the objective can be screwed into the hole with the screw thread tapping its own thread in the plastic. Different body caps have differing plastic density and some are easier than others. If the hole is too loose, then rethread with a fine tape like that used by plumbers to seal water leaks in threads. As long as the lens is not regularly screwed and unscrewed, the objective can remain firmly in place for years. If you have a number of different objectives it is best to use different body caps. The alternative in today’s market is to buy a properly engineered adapter. Many objectives use the Royal Microscopical Society (RMS) thread, but check before ordering. As with other adapters, check availability on auction sites but a good engineering company that produces adapters for most things (and if they do not have it, they may be able to make it for you) is www.srb-photographic.co.uk.

Fig. 3.17 Head of a dung fly, Scatophagus, using a reasonably priced ×10 Joy Optic objective lens on bellows. (×20 magnification, composite image of 48-image stack, diffused flash.) With the objective screwed into a body cap or plate, this can be mounted at the end of a set of extension tubes or on the bellows. Magnification can be increased or

decreased by adjusting the degree of extension. If there is insufficient extension there will be vignetting in the corners. If you need a lower magnification it is best to have several objectives, so a ×4 objective could be used with a greater extension. Working distance will vary but typically a ×10 objective will be between 15mm down to 10mm. The diameter of objectives is quite small at the tip and this allows enough space for lighting the subject. With higher magnification objectives, the working distance becomes appreciably smaller, making focusing and lighting more difficult.

Fig. 3.18 Laowa 25mm f2.8 Ultra Macro lens, released mid-2018, with a range of 2.5×–5× magnification. A strong competitor to the Canon 65mm MPE lens but with manual aperture.

Infinite Objectives During the 1930s, the German microscope manufacturer Reichert developed an idea of changing the objective so that instead of the light rays being focused on exiting the lens, they are parallel. Initially it might seem strange that they do not reach a focus point, especially as this requires an additional lens higher up the microscope to focus the rays to reach the eyepiece. The advantage of this, and picked up and developed more during the 1980s by Leica and Zeiss, allows various optical devices to be

inserted into the space where the rays are parallel, such as beam splitters and polarizers which we come across in Chapter 5. For a diagram of the light pathway, see Fig. 4.10. These infinity objectives will not work with extension tubes or bellows, as the light rays will be in parallel. Instead the objective requires an additional lens to bring the light rays down to the sensor. The technique has been around for some years and is used by the BBC when filming extreme close-ups. The infinity objective is set up on a lens, ideally a 200mm telephoto. Although a prime is best, it can be fitted to the front of a zoom lens set to the 200mm focal length. The adapter is a plate with a hole threaded to take the objective and the round edge threaded to screw into the filter thread of the 200mm lens. The objective needs to be fairly close to the front element of the telephoto and so if this is recessed back inside the barrel of the lens, then it will be unsuited. In the right circumstance the Raynox lenses can be used but, ultimately, try out various options before buying any adapter. A temporary plate can be made up using thick black card. If you do go down the infinity objective route, there are some good Chinese ones available at a very reasonable price; again, like the finite ones, buy if you have the opportunity to exchange them if the quality seems suspect. MACRO AND MICRO OBJECTIVES WITH DIAPHRAGMS During the late 1970s and 80s, camera makers like Minolta, Nikon and Olympus produced specialized bellows lenses which they called macro or micro lenses, derived from objectives. For example, Minolta collaborated with the German microscope firm Leitz. Producing the 12.5mm Minolta Micro Rokkor, it is essentially a Leitz objective complete with an RMS thread and a manual aperture ring. These require an adapter (shown in many of the photos here) to mount on bellows. Nikon produced several, like the excellent Macro Nikkor 19mm, yielding magnifications of up to ×40. In the same mould Zeiss produced a range of Luminars, such as the 25mm one. All can be difficult to find, particularly the Nikkor, occasionally appearing on eBay for silly money. You can be lucky; in 1993 I saw a new 12.5mm in a bargain bin at a second hand camera shop for £25 and snapped it up. Personally, I think the apertures are a waste of time. Stopping down increases distortion, they are best used wide open (or near wide open) with focus stacking to achieve depth of field. Of all of them, the Nikon 19mm would be my favourite: sharp, good working distance and starts where the Canon MPE lens ends. Ultimately, though, they are not entirely perfect and unless available at a good price, there are some equally good ex-microscope objectives out there. Below from the left are the Zeiss Luminar, Macro Nikkor and Minolta Micro lenses.

FINDING THE SUBJECT Photographing extreme close-ups with objective lenses for high magnification definitely requires patience, not least of all when you are trying to set up the specimen and get it in focus. This can be quite frustrating. Start by knowing the approximate working distance so the end of the objective can be positioned close to that point on the subject. Doing this on the horizontal is not so bad and by moving around behind the specimen, you can then reposition it so that it is directly in front of the lens. This can be helped if you are able to watch the image in live-view. On a set-up that is vertical, it can be more difficult and takes some experience. With a DSLR there is a trick of shining a bright light through the optical viewfinder and in dim light, a small spot is projected by the objective on to your subject. This can be taken further by using a laser beam, such as laser pointers. On the forum photomacrography.net, there is a good reference for how to set up your own inexpensive sighting laser that can shine a very sharp beam as a cross. This can be used effectively to focus, even without looking at the live-view. If your photography regularly uses substantial amounts of high magnification, this comes into its own.

FOCUSING RAILS We established early on that autofocus was impossible with extreme close-ups. Even with a macro lens, the best option is to set the approximate magnification for the subject and then move the camera forward and back to attain the focus. However, with depth of field being expressed in microns rather than even millimetres, handholding is virtually impossible except for a few times life-size magnification. You might be able to achieve a reasonably stable base (see Chapter 6) by resting on a table or lying down in the field but to carefully control back and forth movement, especially if you need to focus stack, the only option is to use a focusing rail. Occasionally you see comments made in the photographic press that focusing rails are a waste of time. For what we are trying to achieve in this book, they are so essential, one (or two) should be at the top of your shopping list. Well-known makes such as Manfrotto, Velbon, Novaflex and Kirk are all good but quite expensive. What you are looking for is a rail that can be advanced slowly and smoothly without side deviation. That is quite a demand. By searching online, you will see that there are some very cheap ones to be found and often they come as a pair for around £20. It may seem odd to buy a pair but by attaching one across the other at right angles, it should be possible to help position the camera in two planes rather than just one: backwards and forwards and side to side. This can work well with a good pair and it is amazing how good some are at these low prices.

Fig. 3.19 Infinity objective fitted to the filter thread at the front of an old manual pre-set 200mm lens.

Fig. 3.20 A small selection of manual focusing rails.

Fig. 3.21 Right-angle viewfinders are good in some situations when focus stacking and the rear screen cannot be seen.

Fig. 3.22 StackShot (left) and WeMacro automatic or electronic focusing rails. Cheap does not always mean poor quality, so look at the reviews. One name that is good at this price is the Neewer Pro 4-way focus rail. The rail is on a support and the camera firmly attached to the rail. By winding a knob on the side, the camera can be moved. There will be a locking knob to one side to prevent the rail from moving and after releasing it to advance the camera, you may need to tighten it slightly to cause a small amount of friction and so aid the smooth advance. The length is usually 200mm and may well fit a set of bellows perfectly. The top of the range manual rail is probably the Really Right Stuff B150-B Pkg Macro Focusing Rail, with superb precision movements in two planes. Unfortunately it is expensive at around £500–600, but it is unbeatable quality for a manual rail. As magnifications increase so the care of moving the rail also increases, not to mention the difficulty in managing tiny increments. This is especially so if these are needed in a stack sequence to be approximately equal and where an automatic or electronic focusing rail comes into its own. These use a stepper motor fitted to one end of the rail. As the motor turns, it rotates a screw running the central length of the rail and with guides on either side. A platform with the camera mounted on top is then moved along this rotating screw. A stepper motor is designed for tiny increments of

movement and so the camera can be adjusted by a matter of microns along the rail. So precise is it that a stack sequence can be repeatedly made from the same points, time after time.

Fig. 3.23 Automatic focusing rail control box comparisons. The StackShot control box front (left) and the software window used in WeMacro. Plenty of people make their own as the materials are easily sourced, including the bare stepper motors. Programming the mechanism could be the most complex part, depending on whether you have a tame IT consultant to help out. The amazing Raspberry Pi Barebones computer is ideal for this and some excellent examples of self-programmed automatic rails are out there. There are websites that might help you build your own and help with coding, including www.diyphotography.net. Alternatively you could buy the complete automatic rail system all set up for focus stacking. StackShot has been available since 2011 and is still the leader as it has everything you need for the task. There is a standard 100mm rail and also a 200mm version available. The latter can be the most versatile when working in a studio. The StackShot auto rail has a video cable running to a control box for power and fine control. This box also has a shutter-release cable going to the camera so it can automatically fire the shutter. It is also possible to attach a USB connection to a computer and run the process of stacking from a programme like Helicon Remote. The powerful motor happily supports and drives up to 4.5 kilos of weight, vertically as well as horizontally – an important consideration when using a DSLR, a lens such as the Canon MPE and possibly a macro flash as well. Designed primarily for DSLRs, it can also be used with CSCs along with the correct shutter release cable. The control box can be used to set up a stack or just used to move the camera on the rail for fine focusing with intervals down to 2 microns (two thousandths of a millimetre). Running on 12 volts, it has a mains lead but is easy enough to use in the field with a battery. For seven years, the Cognisys StackShot system from the USA has not only been

the Rolls Royce of automatic rails but really the only contender. During 2016, an excellent alternative to the StackShot became available in the form of the WeMacro rail out of Shanghai. It is a very similar piece of equipment and although physically longer, it still has the same 100mm movement. The finer screw thread allows precision control of travel down to just 1 micron. The control box unlike StackShot has no buttons, just connections and an on/off switch. Linked to a computer by USB, the downloaded software produces an on-screen window of controls the same as the StackShot box, including configuration. Currently, the programme is only available for PCs running Microsoft Windows. (A Mac OS version is due out in the future.)

Fig. 3.24 Empid fly. (×4 magnification, Canon 7D mk2 with 65mm MPE lens on ƒ5.6, twin macro flash, composite of 38-image stack.) However, one advantage of WeMacro is that it can be run from a phone or tablet, both Android and iOS using an app. These use Bluetooth to connect although a possibly more accurate alternative is to use the OTG adapters (they come with the

kit) so that the USB lead will attach to the phone or tablet. The same battery and leads as the StackShot can be used in the field but there is an optional extra in the form of a small battery pack that takes three small lithium rechargable batteries. The most significant feature is the cost, with WeMacro less than half the cost of the StackShot. WeMacro is a fairly new company developing a variety of tools for extreme close-up, from lenses to rails and stands. While still in its relative infancy, changes are ongoing, particularly in software and control. In terms of use, the principles are the same and they are remarkably similar and straightforward to use. We will look at both of these rails in action in Chapter 7. While the function of focusing rails is to move the camera to achieve fine focus on a subject, this can also be accomplished by moving the specimen instead. The heavy camera has a fixed position and the rail can be adapted to hold a specimen that is moved at small increments. With a much lower weight to move, this is an easier and potentially less vibration-inducing option. The drawback is that it is only really viable with dead material. Living material, particularly if it is aquatic, cannot be disturbed. Some very small specimens confined in cavity slides will work. Moving the specimen instead of the camera comes down to so many variables only you can make the decision based on the majority of your subjects, weighing up advantages and disadvantages of both. These issues of support and organization of specimens will be discussed in Chapter 6, but before then we need to see more about the microscope. METHODS AND MAGNIFICATIONS FOR TWO SENSOR SIZES This is intended as a guide only and will vary with equipment, especially different objectives, for example the ×10 finite lens here is a Vickers. The stack interval refers to the distance set between photographs using StackShot for focus stacking.

Methods

APS-C Mag

Full Frame Mag

Approx Working Stack Dist mm Interval mm

150mm macro + ×1.4 extender on 65mm extension tubes

×4.5

–

160

–

Canon 65mm MPE ×1 ƒ5.6

×1.6

×1

95

0.4

×2 ƒ5.6

×3.1

×2

65

0.26

×3 ƒ5.6

×4.7

×3

50

0.16

×4 ƒ5.6

×6.3

×4

44

0.12

×5 ƒ5.6

×8

×5

40

0.09

×10

×6

38

0.05

×5 ƒ5.6 + 60mm extension 50mm coupled to 100mm macro

×3

28

50mm coupled to 300 macro

×7

40

Extension tubes with ×10 objective 13mm extension + adapter

×14

×8

7

0.03

44mm extension + adapter

×20

×12

6

0.02

Bellows with Lietz enlarger lens reversed

Minimum extension

×5

–

40

0.08

×10

–

35

0.04

×7

×4

30

0.1

100mm extension

×10

×6

32

0.05

200mm extension

×15

×9

22

0.03

Minimum extension

×21

×13

4

0.02

100mm extension

×28

×18

4

0.015

×4 Infinity objective on 200mm

×7

×4

12

–

×10 Infinity obective on 200mm

20mm extension Bellows with ×4 objective Minimum extension

Bellows with ×10 objective

×20

×12

12

–

Raynox 505 on 100mm lens

×9

–

12

0.04/ƒ22

Raynox 505 on 200mm lens

×12

–

12

0.04/ƒ22

Chapter 4

An Introduction to Microscopes

erhaps the most obvious optical device for enlarging subjects to either view or take photographs is the microscope. The instruments can be quite expensive to buy and complex to use. Over many years they have evolved from basic lenses into elaborate designs where entire modular systems can take a multitude of filters, different lenses and illumination sources. This chapter is intended to provide an introduction to the device, what to look for, how to use it and most importantly where to find out more. While they provide superb ways of photographing material beyond the realms of the previous chapter and are very satisfying to use, they are complicated; if you take on the challenge expect to be engaged in a major project that could take over your life! Most books dedicated to the subject are not particularly for the faint hearted and hence this introduction. It cannot be emphasized enough that if you decide this is a project for you, consider joining a group – but more about that later.

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Fig. 4.1 Forty-nine diatom exhibition slide arranged and produced by Klaus Kemp. Photographed with dark field illumination on a Wild M20 compound microscope using Sony A6500 body. (Magnification approximately ×80.)

TYPES OF MICROSCOPE AND THE PROBLEM OF THE STEREO

There are two types of optical microscope: stereo and compound microscopes. The latter is probably what most people will think of as a microscope.

Fig. 4.2 Beck stereomicroscope, circa 1930s. The principle of the stereo, like two microscopes side-by-side, can be seen with the two eyepieces and two objective lenses. Already we have looked at objective lenses on their own as a way of magnifying subjects. This is located at the bottom of a tube and to pick up the image passing

through the tube is an eyepiece (or ocular) at the top to look down. In essence, that is the optical make-up of a compound microscope. To make looking through the eyepiece a little easier, some microscopes have a binocular head (an eyepiece for each eye) to reduce eyestrain when using it for long periods. Stereos provide a threedimensional view of the subject and compound microscopes produce a rather flat, two-dimensional image. That is where a stereomicroscope comes in, as they do provide a 3D view. To do this, they are like two compound microscopes held side-byside with one for each eye. As our eyesight and brain are set up for this stereoscopic perspective, this type of microscope produces a clear and detailed view. Good stereo imaging, however, is limited to lower magnifications for optical reasons and all stereomicroscopes are generally in the range of around ×10 to ×60.

Fig. 4.3 Modern stereomicroscope with the third tube for taking a camera, although the image quality is quite limited. For our purposes this may seem fine, but there are major downsides. High optical quality comes at a price and new stereomicroscopes with names like Leica, Nikon and Olympus will be very expensive indeed. Secondly, the low and average priced stereos do not make high-quality photographs. When looking to buy there will be examples fitted with a third tube (called a trinocular as it has three oculars or eyepieces), specifically to take a camera. This can be a DSLR or CSC although

some makes have a dedicated ‘block’ type. It really does not matter as unless the optics are of the highest quality, none will produce good, sharp images. Figs 4.3 and 4.4 show two stereomicroscopes designed with photography in mind.

Fig. 4.4 Wild M3Z model stereomicroscope from the 1980s specifically designed for photography. The first is a recent GX system at around £400–600 that makes an excellent instrument for viewing and zooming. LED lighting can come from within the base

(transmitted) or from a spotlight above the stage for top lighting. As a mechanism for taking photographs it is not recommended, as quality is limited. The second stereomicroscope is the Swiss-made Wild M3Z. Wild was taken over by Leica in 1994 and continued for a while with both names. The M3 series of stereos were produced during the 1970s and 1980s and were one of their most successful and high-quality optics. The substantial third tube easily takes a DSLR but the left eyepiece is also designed for photography. A lever on the side centres the optics and blanks the right eyepiece so that a camera attached to the left eyepiece will have an optimized performance. The M3Z is set up for very controlled, transmitted light coming through the base and two stanchions in each back corner will hold lighting for top, reflected light. M3 stereomicroscopes can be purchased second hand and are always available on auction sites at prices starting at those of the GX system. The M3Z will be substantially more as the M3 stereos were developed as a system and parts are available either second hand, new or from a third party like Motec. In this way you could start with a basic system and buy photo tubes for cameras and different stages. Both the GX and the M3Z shown here have a magnification zoom, but this is not always the case. Lenses are not quite as flexible to swap as compound microscopes although Wild did produce an excellent Planapo objective for the M3Z. This might be enough to put you off a stereo but they are useful in preparing material for use with your various photo techniques. Buying one for just a few hundred pounds will provide you with an ideal ‘dissection’ stereomicroscope. A large flat base where specimens can be placed to view under the objective is ideal. Illumination will be by either fitted or separate lights that shine down on to the material. A stereomicroscope with a boxy base will have the lighting inside and transmitted up through a translucent base; ideal for small glass dishes with aquatic creatures. That will help to show up transparent organisms so they can be caught with pipettes and transferred to other containers to photograph on other equipment. A stereomicroscope is a useful piece of equipment to help view and prepare material for extreme close-up photography. Buying one for photography would require a substantial outlay of money, even for a second hand system. However, there may be another option to consider.

USB AND INSPECTION MICROSCOPES These inspection microscopes were mentioned with block cameras and, along with USB microscopes, are a fairly recent development. Both types can connect to a computer where the image is relayed to the screen for viewing. Magnification is around that of a stereomicroscope starting from 1:1 up to 20:1 or higher. The cheaper USB-only versions produce low-resolution images of 0.5 up to 2 megapixels

at the most. This could be used as a record shot, demonstration and for emailing but not so good for general photography.

Fig. 4.5 Industrial inspection HDMI microscope digital camera suitable for 12MP and HD video with magnification of ×1–×25.

Fig. 4.6 Close-up of the camera controls on the top of the HDMI microscope. The inspection microscopes are a different breed altogether, remarkably good value and well worth looking at. They do not have the sensor size to provide exceptional images but they produce quick and easy photos as long as there is a careful use of lighting. The main problem is the name. Inspection microscope covers such a diversity of products that it can be difficult to narrow down the type of microscope we are after. Searching for them online normally requires a description like ‘HDMI HD Microscope USB Industrial Camera’. This should bring up fairly

complete systems like the one shown in Fig. 4.5. The basic camera and lens will be just over £100, but it can be worth buying it with a sturdy base, LED lighting and remote control. The camera unit needs to be around a 10–14 megapixel unit and will be a square block which screws into the microscope column with a zoom objective at the end. On the top of the camera are push buttons, which control exposure and shutter release. There is a slot for a micro-SD card and sockets for the power supply, a USB link to a computer and an HDMI socket that can connect the camera to a monitor or TV output. Supplied software can be installed on the computer so that when the USB link is made, the image can be seen on the screen and photos taken. These are usually low resolution, around 1.5 to 2 megapixel images maximum despite a higher pixel sensor.

Fig. 4.7 Industrial inspection microscope with HDMI connection to a 4K monitor giving a live feed. There is a top LED light and additional Ikea Jansjo spots to illuminate the moth. Before installing the software, there may already be suitable applications available

on the computer. For example, Windows 10 has a camera app which when opened usually defaults to the connected USB camera. Minor adjustments can be made and photos taken by clicking the screen and the image is then stored on the computer. The camera does not allow two different simultaneous links and the HDMI connection is the best option. An HDMI cable will produce a good image on a computer monitor and then using either the remote or the controls on the camera, photographs can be taken. These will utilize the full sensor area to create high-resolution images and save them on the micro-SD card. The pictures can be reviewed on the screen and either downloaded via the USB cable or by removing the card and transferring to a computer.

Fig. 4.8 Dog flea on a glass slide and with back illumination. Photographed using the HDMI microscope. (×20 magnification.) Like any microscope, lighting is key and with the right set-up, remarkably good photos can be achieved. The fact that the image is blown up on the monitor means that focusing and stacking is straightforward. Like the stereomicroscope, perhaps one of the most useful features of having an inspection microscope is in helping you to sort through sample material to find and orientate subjects ready to photograph with other techniques. For example, a dish of small seaweeds in seawater from a rock pool full of creatures that can be picked up and transferred to other dishes to

photograph with a DSLR or higher magnification microscope. Along the way, before the transfer, you can take record shots.

Fig. 4.9 Stereos and HDMI microscope comparison using part of a sea urchin test magnified approx. ×5 all with similar LED lights. A: a GXVision stereomicroscope (Sony A6500 camera body); B: Wild M3Z (Sony A6500 camera body); and C: HDMI microscope.

INTRODUCING THE COMPOUND MICROSCOPE

The objective is the lens close to the subject, which relays a real image to the eyepiece. This real image is then magnified by the eyepiece to create a virtual one which we see. Both of these compound lenses will have their magnification written on them and to work out the total magnification of the system, multiply the two together. So a ×4 objective with ×10 eyepiece gives a total of ×40 magnification.

Fig. 4.10 Diagrammatic views of the light path through the compound microscope. A: with finite objective lens; and B: using an infinite objective. The eyepiece is important to project the virtual image on to our retina but in photomicrography, the eyepiece can be replaced by a monitor or camera sensor. In the case of the infinite objectives introduced in the previous chapter, the rays of light exiting the lens are parallel and so there is an additional lens inside the tube that brings the focus down to form the real image within the eyepiece. The term ‘field of view’ refers to the area of the object that is visible down the microscope. This is determined by the objective lens magnification and is further enlarged when it is

projected on to the eye. This image is circular and our eyes can record this, giving us the full field of view. As sensors in cameras are rectangular, the image will be a smaller, cropped part of the field of view.

Fig. 4.11 Second hand compound microscopes. A: a basic and reasonably priced Vickers with bright field illumination. B: Wild M20 trinocular, ideal for photography. There are various designs (which over the decades have changed considerably) for supporting the microscope tube. Additionally this tube has to be moved smoothly up and down to focus on the specimen located on a stage. The process of focusing is done first by a coarse focusing knob that achieves approximate focus, before adjusting with the fine focus. The specimen is placed on a microscope glass slide, not directly on to the stage. Ideally the slide is moved by a mechanical stage fitted to the basic stage. By rotating two knobs on the side, the slide can be moved up and down and side to side in a precise fashion. Basic models will be missing a mechanical stage

but, with care and experience, at low magnifications it is quite possible to do this by hand. When beginning to focus, do not look down the eyepiece but instead look from the side and using the coarse focus, bring the tip of the objective down towards and near the subject. Then looking through the eyepiece, the objective can be raised away from the specimen. By focusing away from the specimen, you avoid touching (and crushing) it with the objective. Perhaps the most important factor after the lenses is what is located under the stage. A hole in the latter allows light to pass through to the specimen. In most cases, compound microscopes utilize transmitted light, coming from below the specimen. Due to the closeness that most objectives must get to the specimen, reflected light from above is difficult. We will look at this separately as it requires a special set-up called epi-lighting (which will be covered in the next chapter). A very basic microscope will have a mirror beneath the stage to reflect light up through to the glass slide. This is not easily controllable and most microscopes use artificial light either from a tungsten, halogen or LED light source.

Fig. 4.12 Three different sub-stage condensers that fit the Wild M20 microscope viewed from the top. The cap in the foreground is a flip-out top lens that needs to be fitted on the condenser top for high magnifications. The condenser on the left is a universal one that rotates to change the type of lighting such as bright field, dark field and phase contrast. Between the mirror or light and the stage should be another structure. In basic microscopes this could be a rotating disc with various sized holes to control the amount of light passing from the mirror or light. A better arrangement is a small tube with a lens at the top and iris diaphragm at the bottom. The lens directs the light whilst the iris controls the brightness. This tube is a simple type of sub-stage condenser. As Fig. 4.12 shows, they can be quite variable in size and complexity, important for changing the lighting and for adding filters and discs into the beam of light. Even illumination of the specimen is very important. If the lighting is not set up correctly, a quick look through the eyepiece will reveal a bright area in the middle, grading to darkness at the edge. If the light is too strong and not focused correctly, a specimen loses contrast. This is the job of the sub-stage condenser. Most microscopes can be fitted with different sub-stage condensers to accompany

particular objectives and magnifications. Some condensers are fitted with a flip-out top lens, to ensure full illumination of the field of view for low power objectives. The condenser is located and fixed in a holder that can be racked up and down to focus the light correctly on the specimen. The technique used to set this up correctly on all modern microscopes is called Köhler illumination.

KÖHLER ILLUMINATION This method of adjustment to the lighting will provide the best conditions to achieve an accurate image and will not be possible on very basic microscopes lacking sub-stage condensers. You will need a specimen on a slide, preferably a permanent, prepared slide such as a stained section of plant material or translucent insect parts; something that has good contrast and translucency. Turn on the microscope light and open the field diaphragm to allow plenty of light through. This iris diaphragm is not always obvious, located beneath the condenser and maybe on the lamp housing. As an example, Zeiss microscopes have the lamp in the base and so the iris is where the light emerges from the base. On the Wild M20, the iris is in the light housing as it enters the base. Place the specimen on the stage, select the ×10 objective (making sure it has clicked into place) and adjust the focusing controls, as outlined above, until the image is as sharp as possible. Now close the field diaphragm completely. As you look through the eyepiece, the field of view has disappeared to a bright spot of the subject in the centre. The edge of this spot is the out-of-focus and blurry edge of the field diaphragm. This needs to be in focus and to do that, the condenser is raised or lowered. Use the knob on the side of the condenser holder to do this until the edge of the spot becomes sharp. That shows the field diaphragm is projected precisely on to the specimen. The field iris can now be opened to widen the field of light until it is just beyond the edge of the field of view. Your view should now be evenly lit but not so that there is too much stray light entering from the side and reducing the contrast of your image.

Fig. 4.13 Setting up Köhler illumination. A: switches on/off and controls the amount of light; B: field diaphragm; C: raises and lowers the condenser; D: condenser diaphragm; F: the filter holder; MS: the mechanical stage. There is another iris diaphragm below the stage, located in the base of the condenser, which controls the amount of light entering the microscope. As much as anything, this affects the contrast of the image. Too much light reduces contrast while closing the iris to approximately two-thirds of the field of view helps to increase the contrast (closing it more than this will introduce diffraction into the image). Although

this form of lighting is referred to as bright field illumination, the procedure is a good starting point to set up all microscope lighting and needs to be repeated for different objectives. In some cases, a flip-top lens may need to be added to a condenser but really for any change in objective, the Köhler method needs to be employed to reach the most accurate lighting and then adjust the condenser iris as well.

TAKING PHOTOGRAPHS THROUGH A MICROSCOPE Cameras with a fixed lens can work as long as the diameter of the lens is sufficiently small to fit over the eyepiece. This will still require the use of a zoom to minimize the vignetting effect. Quite simply, replace the eye with a compact camera or smartphone by placing the lens over the eyepiece and looking at the rear screen. To minimize vignetting as much as possible, try zooming in to a telephoto; this increases magnification and reduces vignetting as it goes. Fig. 4.14 was taken in this way, supported in place by a tripod, although it was held by hand as the shutter was released at around one thirtieth of a second. Depth of field was so limited that three photos were taken, each time shifting the fine focus on the microscope slightly to focus on a different part of the anatomy. The images were later combined on the computer. Vignetting is always a problem, even with zooming in on the specimen, but this can be improved on the computer in a program such as Adobe Lightroom. Tripods are not very practical for regular use in this way and various platforms or clamps can be purchased. Always get the specimen in focus first and then add the camera, which then autofocuses on the virtual image coming out of the eyepiece. Sometimes switching the compact to close-up function can assist in achieving focus. This method works especially well on basic microscopes, which only have bright field with limited controls.

Fig. 4.14 Head of a phantom midge, Chaoborus, larva. Common in ponds, it is a predator of water fleas, catching them with the modified antennae. (×40 using bright field illumination.) Taken on a point-and-shoot camera, the lens was held over the eyepiece to take three images that were later stacked to form this composite example. Postprocessing on the computer removed the vignetting that caused a darkened gradation to the corners. The specimen was live in water on a cavity microscope slide covered in a cover slip. (Sony compact on Chinese student monocular microscope.)

Fig. 4.15 The photograph in Fig. 4.14 was taken by holding the camera lens to the eyepiece. When using an interchangeable lens camera, the lens needs to be removed from the body so that an adapter can be fitted. The one shown in Fig. 4.16 is a universal adapter from a third party, purchased from eBay. The diameter of the microscope tube needs to be checked so that the adapter tube slides in easily. An eyepiece from the microscope can be removed to insert the adapter or better still with a trinocular, the camera can be permanently mounted on the third ocular tube. Adapters specific to the maker of the microscope can be a better bet if they are obtained at a reasonable price. The camera body is attached to the specific tube adapter by a T2 mount, easily obtained online.

Fig. 4.16 A DSLR adapter to fit the camera body into the tube of the microscope. Note the lens at the end of the adapter, which replaces the eyepiece, to project the virtual image on to the sensor. The main advantage of the correct make of tube over the universal adapter is in maintaining the working distance of the objectives. This is the distance from the tip of the objective to the specimen when the virtual image is viewed in focus through the eyepiece. As the length of the microscope tube is standard, this distance will be designed for particular objectives so that they are all in focus (parfocal) when changed from one to another. The problem comes if you use an adapter that changes the distance from the objective to the camera sensor and the microscope has to be refocused, as the working distance will change. What you see in focus through the eyepiece will be out of focus in the camera, and vice versa. In fact it is quite easy not to achieve a parfocal set-up as the tube length only has to vary very slightly and the image on the sensor will not be sharp. Slightly raising the eyepiece on a small collar or cardboard spacer may help. Live-view will help with this as you look through the microscope and check the camera view, separately blown up on a screen.

When viewing specimens, we have already mentioned using a glass microscope slide to support and move them. In most cases it is best to cover them too, especially if it is in a liquid such as aquatic plankton. For this you need a cover slip, which also prevents contamination of the objective. Both the cover glass and slide will change the way that light will refract through to the objective. To maximize the image quality, objectives are corrected to cover a range of thicknesses and this is denoted on the lens, often from zero with no cover slip, through to 0.17mm thick.

BUYING A COMPOUND MICROSCOPE Microscope shops, if they even exist, are few and far between. Online there are several companies, such as Brunel Microscopes, which can assist in finding a good model. Auction sites are good if you know what you are looking for. Ultimately, the best option is to join a group such as the Quekett Microscopical Club before buying a microscope. At the numerous meetings held around the UK, you will meet wonderfully supportive and helpful members. They will be able to recommend and discuss equipment and invariably know someone who is selling good items at reasonable rates. They will also provide assistance in setting up and using the microscope. Several times a year members bring their spare equipment along and sell everything from odd and rare objective lenses to complete microscope systems. Details of groups are given in the References section.

Fig. 4.17 Tintinnid from a marine plankton sample. (×100 magnification.) They are very common in coastal seas but have a low survival rate once caught and so the photograph has to be taken quickly. They are so transparent that to make the tube in which they live stand out, phase contrast lighting was used. (Wild M20 microscope and adapter shown in Fig. 4.16.) One starting point could be to work out a budget. There is a bewildering array of microscopes from £100 to many thousands: £100 will buy a new Chinese student monocular microscope with a mirror for bright field illumination and three objective lenses. Adding a binocular head, mechanical stage and a condenser will at least triple the cost. As well as the Chinese manufactured examples, there is quite a collection of student and research microscopes available new from India. India has an established history in microscope building, although for many years the optics came from China. Research microscopes will be either binocular or trinocular, with better condensers and built-in lighting. There will be provision (although not always included) for dark field, polarized light and other illumination systems to be discussed in the next chapter. Possibly the recommended route would be to look at second hand microscopes with a good brand name like Reichert, Vickers, Wild, Zeiss, Leica, Nikon, Leitz and Olympus. Some may have had a hard life in a laboratory, with the accompanying wear and tear. At the same time these may have been regularly serviced and would be still worth considering. As most of the research microscopes will be part of a system, there will not be just ‘a model’ but multiple variations. Look first at the overall state of the bare bones of the frame and smooth running of the focusing knobs. Then

it is down to the lenses; what objectives are sold with the microscope, considering features such as magnification and quality; plan lenses will be more expensive. Wide field (WF) eyepieces are useful over the standard ones and make sure the condenser is suitable for your needs. All lenses need checking for clarity and dirt by looking through them carefully. Objectives can be damaged at the tip of the lens where they have hit specimens or solvents affected the cement holding the glass in place. Buying a ‘system’ type microscope means that lenses you want can be added at a later date. During the 1960s and 1970s, some excellent models were made with high-quality lenses and are available at sensible prices, running into hundreds of pounds rather than thousands. The Wild M20 in Fig. 4.11 is an example of a good model from the 1960s, fairly easily sourced, plenty of parts available with bright field, dark field and phase contrast illumination and can be found for a range of £400–600. There are many good Zeiss examples including several Photomic models from the 1960s. The Zeiss models can be large and weighty microscopes but are easily adapted for digital photography with LED lighting and suitable cameras.

Fig. 4.18 Bosmina, a species of water flea common in lakes in the spring; ×80 using phase contrast lighting to increase the contrast of the live specimen that was in water on a cavity slide. To slow the activity of the creature, a watered-down wallpaper paste had been added. (Wild M20 microscope and adapter shown in Fig. 4.16.) More recent systems from the 1980s include the excellent Olympus BH-2 or Leitz Ortholux. These are just a few examples of the microscopes available. This chapter has merely touched the tip of the microscopy iceberg and hopefully it has whetted your appetite to go further. Microscopes, as with all aspects of extreme close-up

photography, are very dependent upon lighting – and that is the direction we need to take now.

Chapter 5

Lighting Extreme Close-ups

s any photographer knows, the light will make or break an image. For landscapes, the light is best early or late in the day when the sun is low in the sky to cast shadows, often long, dark lines, to contrast with brighter sunny ones. Shadow provides relief to create a three-dimensional image that would otherwise be flattened by the sun at midday as it removes the shadow. An overcast sky diffuses the light, creating softness but rather a flat subject. All of this is down to the fact that light travels in straight lines, being both ideal and a hindrance for photographers depending on the subject. With no cloud in the sky, the straight rays of sunlight skimming across the landscape hit solid obstacles like trees and walls so that shadows occur. This shadow is not completely black as light will bounce off objects and be reflected to these darker patches. When the sun’s rays enter cloud they too start to bounce off the water molecules and the thicker the cloud, the more they are diffracted. By the time they have passed through a large expanse of cloudy sky, the rays are approaching the subject from many angles as it is so diffused. Now the shadows are no longer strongly defined.

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Fig. 5.1 Stinging nettle, Urtica dioica, a close-up of the sting cells. A single, directional flash from the side was used to make the cells stand out against a black background. When attempted with diffuse light, the sting cells did not appear so sharp. (×7 magnification, Canon 65mm MPE single flash from twin macro flash. Composite of 52-image stack at ƒ5.6.) This emphasizes the important principles when organizing light for extreme close-up photography, whether you require shadow and straight rays of light or a softer, more diffused light. Whatever light is required for taking the photographs, you will have to

supply the source because the closer to the subject and the higher the magnification, the less light there will be. The question is not do you need to illuminate your specimen, it is what light are you applying and how? The chances are that using natural light is not an option.

USING ELECTRONIC FLASH A flashgun has so many advantages. Light can be delivered directly to where it is needed and it will be bright. Even the most basic flash will be more powerful than a spotlight and so a large flashgun is unnecessary. Cheap and cheerful is fine. The colour of electronic flash is corrected for daylight and rarely will the white balance of the photo require retouching. Flashguns work like a strobe, that is, rather than firing off one burst of light when you release the shutter, modern equipment triggers a rapid sequence of short flashes, each of thousandths of a second. These individual flashes build up until sufficient light has illuminated the subject and then the strobe is switched off. To us, it appears as one quick flash. Synchronized with a moderate shutter speed, even if there is some vibration of the equipment or specimen, the short flash duration should prevent subject blur.

Fig. 5.2 Young adult wasp emerging from its pupal cell in the nest with a larva below. Live specimens photographed in the field. This is a composite of a sixteen-image stack using twin flashes to freeze any movement but there is reflection from damp, shiny areas around the head as the light is too harsh and needed diffusion. The composite needed retouching as both adult and larva moved during the stack. Flash can be used in most situations, even under microscopes. One possible downside is their direct bright intensity that can produce harsh shadow and strong reflections on shiny subjects. By employing diffusers, these problems can be

controlled and eliminated. Another issue to remember is the recycling times of the flash, the time taken for the capacitors to recharge in time for the next flash. In most cases this is not a problem as working up very close to a subject rarely causes the complete discharge of the flash. You will see that the charge light remains on. Problems can develop during a recycling times of the flash, the time taken for the capacitors to recharge in time for the next flash. In most cases this is not a problem as working up very close to a subject rarely causes the complete discharge of the flash. You will see that the charge light remains on. Problems can develop during a recycling times of the flash, the time taken for the capacitors to recharge in time for the next flash. In most cases this is not a problem as working up very close to a subject rarely causes the complete discharge of the flash. You will see that the charge light remains on. Problems can develop during a long focus stacking sequence where one photo is taken shortly after another for a prolonged period. This is not necessarily down to a recharge time but an inherent issue with capacitors. For example, we are going to run a hundred image stack on an electronic rail. More detail will be discussed later on, but try to keep with the scenario. A two-second gap is set between shutter releases for both the recharging of the flash and a settling down period to stop vibration. However, halfway through the stack, the camera and flash can suddenly refuse to fire when required. After a refusal of around eight to ten seconds, the camera fires again. This may happen repeatedly and is down to the flash and certain cameras. For example, some Canon bodies can be prone to it when using a basic flash. Also this is not down to the batteries or whether the flash is wired or wirelessly connected. To get around the problem, if it occurs, just increase the time interval between shutter releases – try a gap of four to six seconds. Flashguns designed for macro and close-up work, such as a twin macro flash, should have negligible problems if any. More about these flashguns below.

Fig. 5.3 Lesser water boatman, Corixa, live specimen in water in an observation cell, behind glass, using a twin flash set-up with the main, brighter one on the right and a fill-in, dimmer one on the left just to take the edge off the shadow. Note both flashes were kept at around a 45-degree angle to the glass so no reflection occurred. (×3 magnification, composite image from a 42-image stack.) Always make sure that fresh rechargeable batteries are put in the flash before a long session of focus stacking, as it can be difficult as well as annoying to change batteries part-way through. Ordinary rechargeable batteries discharge themselves over a few weeks. Eneloop batteries are by far the best type to use as these lose charge only a fraction over a year or more. These reliable batteries were originally produced by Sanyo, but are now available from Panasonic and only cost around 20 per cent more than ordinary rechargable batteries.

FLASH DIRECTION AND THE NEED FOR

DIFFUSION Flash is therefore a good and flexible way of lighting extreme close-up shots, but how should this be positioned? The subject matter will determine what light is best and we will look at specific scenarios later. Always have a quick analysis of the specimen with a hand lens so the nature of the material can be assessed. Does the subject have areas of shiny, reflective surface? Small nooks and crannies that need shadow to give 3D relief? Dense hairs can be tricky as they could merge into one apparent lump rather than stand out individually and transparent areas can disappear completely. Flash may be ideal, but the resulting image depends on its careful use. In nature, light normally falls from above. You need to bear this in mind when photographing aquatic organisms as lighting from below can look unnatural. Remembering that light moves in straight lines, use a flash against glass by holding the flashgun at 45 degrees to the glass surface. As the rays hit the glass, some pass through but the majority will be reflected off at 45 degrees. If the camera lens is in front of the glass, then the reflected light will not enter the lens. The problem with a flash mounted on top of the camera, along the axis of the lens, is that the rays hit the glass at 90 degrees and bounce back at 90 degrees, straight back into the camera. Ideally, then, try to hold the flash above the specimen and at a 45-degree angle for the light to be more natural and lack reflection.

Fig. 5.4 Wing of a clouded yellow butterfly photographed with a ×10 objective lens and one direct flash held on the right skimming light over the surface to create shadows on the left side of each scale. (×22 magnification, composite image from a 58-image stack.) With no glass to be concerned with, the angle of holding a flash, although more flexible, is still useful at an oblique angle. Like the sun at midday, if the flash shines directly on to the surface, shadow is minimized along with the texture. Fig. 5.4 shows the use of a single flash held almost flat to the butterfly wing so that directional light skims across the surface of the scales, forming slight shadow on the other side. Just

like a landscape with the sun low in the sky, the image has more texture. This photograph was taken with a Vickers ×10 objective on a bellows unit and the working distance was around 4mm. In this instance one flash worked well, but this is not always the case. The problem with one flash is that it can create strong shadows and this may need to be tempered by a second light source on the shadow side to tone this down. If the intensity of the second light is the same then the shadow can be removed altogether, but by varying the amount of light, a harsh shadow is reduced to a slight presence just to create enough texture in the photo. This fill-in light could be a second flash or a reflector. Small reflectors can be made out of little loops of wire with silver foil glued to them and then are positioned close to the opposite side of the main flash. It may require several test shots to get the reflector in the correct position. Reducing the intensity of a second flash may require a few layers of white cloth or tissue fixed over it. An alternative is to make the main flash more intense and directional. For many years, fashion photographers have used snoots, cones that narrow down the light like a funnel, to produce a narrow beam. These are easy enough to make, especially for extreme close-ups as everything is on a much smaller scale. Essentially any suitable rigid cardboard tube can be adapted, or a section of a cereal packet rolled up into a cone. Strength and shape can be maintained by gaffer tape, which can also be used to hold it in place on the flash. The cones that hold yarns of wool to fit on a knitting machine are perfect if you have a chance to obtain them. The inner surface of your cone – as well as diffusing the light – might tint the colour. Lining the cone with kitchen foil along the sides will make it more reflective, as well as preventing tinting. The end of the cone snoot can be left open or small, clear plastic lenses can be sourced cheaply to further direct the light. A snoot on a flashgun was used in photographing the butterfly scales in Fig. 5.4, as the working distance was so narrow and it was easier to control the direction of light.

Fig. 5.5 Minolta ring light. Originally designed for film, it can be modified for digital by connecting it to the camera via a voltage regulator called a Safe Sync. This reduces the trigger voltage to protect the camera circuits. Another way to intensify the light is to use a bunch of narrow tubes over the flash. This can be done using white drinking straws. Cut them to a similar length and wrap them up in gaffer tape to attach to the flash gun. Although this works well for macro, the snoot is better for higher magnifications.

Fig. 5.6 Here, a pair of twin macro flashes have been placed either side of a specimen to provide side light. A disposable plastic food bowl is upside down over the specimen to diffuse the flash and the hole in the base allows the photograph to be taken with the camera held vertically above. If you already have a flashgun you may think of just buying a cheap, low-powered flash as a fill-in. Meike manufacture good small flashguns that are widely available on eBay and other sites. Two can be purchased and linked by a commander unit to be paired up so they work together. Have gaffer tape available so that white fabric, such

as bed sheeting, can be fixed across the flash to reduce intensity. The ultimate situation is to have a macro flash unit with twin flashes such as those produced by Canon and Nikon. The power of each flash can be controlled so that a ratio of light of one to the other can be set. If you are operating the camera through a utility on the computer and live-view, the flash can be controlled here without the need to touch the flash. The flash units connect wirelessly to the Nikon commander whilst the Canon flashes are wired. The latter may seem old technology but the advantage is that the flashes obtain their power from the rechargable batteries in the commander. The Nikon flash units have separate batteries, which may not be locally obtainable so if you use these, make sure you keep plenty of spares. It can be surprisingly demanding on batteries for long stack sequences. The biggest drawback to using dedicated macro flash is the very high cost of buying them in the first place. Typically, photographers buy the cheaper option of a ring flash. Low cost versions have a continuous ring of light but more expensive ones will be at least divided into a left and right light that can be adjusted separately. These are nowhere near as flexible as a twin macro flash where the separate flash units can be removed from the end of the lens and fixed to brackets near the specimen. An alternative to the manufacturer’s own twin flash is that produced by the Chinese firm Yongnuo called the YN-24EX Macro Twin Lite, available at a sixth of the cost but not as flexible as the Canon and Nikon units.

FLASH DIFFUSERS Many instances call for less intense, soft and diffuse light. Any subject with reflective surfaces will cause haloes and bright, burnt-out regions. The nettle stings in Fig. 5.1 are very close to being intrusive but the problem was that with diffuse light, the glasslike cells would not stand out sufficiently. Insect exoskeletons are often shiny and there is a play-off between vivid, sharp colours with a bright flash and the muted conditions produced by a diffused flash. My favourite diffuser has always been a ping-pong ball with a hole cut in the surface so that the insect can be placed inside. The flashes are positioned so that they illuminate the insect through the ball material and not the hole. To hold the ball in place, it is glued on to a small wooden dowel that can be held in a clamp and manoeuvred. There are a surprising number of disposable items in the home suitable for creating diffusers, especially those that can be placed over the subject like a tent. Many desserts come from supermarkets in fairly thick individual bowls. As long as they do not have writing on the side and are not coloured (otherwise the light can be tinted as it passes through), the base can be cut out and placed upside down over a specimen. The camera then shoots through the base while the flash units are on the outside of

the bowl. That works if the camera is mounted vertically. If it is horizontal, the cut base can be stuck to a ring that fits on to the front of the lens with the flash on either side. Even attached to a bracket, the important point is that it needs to be held in place between the flash and the specimen to diffuse the light. One reason the inverted dessert bowl concept works well is because the flash is not too close and it has a thick, curved surface. Diffusers flat to the flash surface will produce some diffusion but do not eliminate ‘hotspots’ – bright reflective patches on the specimen. These bowls can be fixed to the flash head either by Blu Tack or gaffer tape (see flash in Fig. 3.10). To increase diffusion further, stuff a small amount of crinkled-up kitchen greaseproof paper or tissue in the bottom of the bowl. This is worth trying if small hotspots appear. Instead of clip-on plastic diffusers, it is possible to buy ‘softbox’ attachments. Although they are not suitable for extreme close-up photography the principles can be utilized. The most notable feature is that they are large and the main diffuser part is some distance from the flash. This can be replicated and tested by making your own diffusers from mouldable plastic. Polymorph is one of several makes of mouldable plastic that can be purchased online. Put a small amount of the granules into a large glass bowl and add boiling water. Within a minute or so the pellets go clear and coalesce. Make sure they have gone completely clear before using an implement like a spoon to take what is now a lump out, drain and place on a chopping board. Using a rolling pin, it can be rolled out into a rough sheet. After fifteen to thirty seconds, as it becomes too difficult to continue, pick it up in your hands and begin, using your thumbs, to push the edges out to increase the size. If the sheet folds onto itself the sides will stick to each other, but not to you or the implements. At any time drop it back into the hot water to clear and soften again. This can be done any amount of times, but once it is in your hand and cooling, the plastic starts to go opaque. At this point drop it over a small, upsidedown glass tumbler and wrap the plastic around the glass, pressing the plastic together to take on the form of the mould. The plastic easily comes off glass moulds and even cones of thick cardboard. Using shearing scissors, trim the edge. If the plastic has hardened, hold the plastic with the edges back in the hot water to soften. When soft, dry with kitchen roll and try to mould the edges around the flash as well as possible. Elastic bands, clips, gaffer tape, Blu Tack or any number of other ways can be used to later hold this in place. If the diffuser does not work out right, just place it back in the bowl and add boiling water to make it soft enough to mould again.

Fig. 5.7 Mouth of a gold-ring dragonfly, Cordulegaster boltonii. The opening is guarded by numerous hairs and shiny reflective surfaces. Two flash units on either side were strongly diffused to prevent reflection and small silver reflectors were placed just out of shot top and bottom to bring light into the mouth. (×9 Canon 65mm MPE with extension tubes and macro flash; ƒ5.6 composite of 56-image stack at ƒ6.7.)

Fig. 5.8 Head of a small click beetle, Elater pomonae, photographed inside a ping-pong ball for diffusion so that the golden hairs and pitted exoskeleton of the head and thorax could be seen. (×8 magnification, dead specimen, composite of 68-image stack, Canon 65mm MPE with extension tubes and twin macro flash.) Diffusers of any shape or size can be made this way, tried out and then remoulded as necessary. Also if the diffuser area is not thick enough, just add more material after softening it up in hot water. Keep all the pieces – no matter how small – that you may cut off, to use at a later date. It is magic stuff and its use is only limited by your imagination. You can never have enough diffusers! As well as the above, it can be worth making up some paddle diffusers. There are times when there just is not enough working distance to organize the lighting and the larger diffusers. These paddles can be of quite variable sizes. Using wire lengths (garden wire works well), bend them into ovoid-shaped loops. Cut out pieces of greaseproof paper, plastic from milk bottles or mouldable sheets that more than cover the loop and trim so that they overlap a little. Then glue them on with Araldite or Gorilla Glue. Make sure you leave plenty of wire coming from both ends of the loop; this can be bent to form legs or wrap around

small stands like ‘helping hands’ (Chapter 6). Paddles are especially useful when using continuous light.

Fig. 5.9 Diffusers. Those for the top flash are made of Polymorph mouldable plastic. The left twin macro flash has a Sto-Fen diffuser fitted. The right flash head has small metal pieces glued to the flash so that varying diffuser material can be attached with small magnets. DIFFUSERS ON FLASH UNITS There are plenty of manufacturers making plastic diffusers that clip on to a flash. Sto-Fen is one such make. On the whole, they are a waste of money as they are quite expensive and after all, they are just a piece of white plastic. With limited use they can be effective on some macro material, particularly out in the field. I have made any number of different diffusers to fit my macro flashes and I start by gluing small pieces of steel on either side of the flash. These are actually the ends of cheap craft knives where, as the end of the blade blunts, they can be snapped off with pliers to reveal a new, sharp end. I then cut strips of plastic from items such as milk bottles and hold them in place (sometimes several layers) with small, powerful magnets. These are easily bought on eBay but I get mine from the bottom of Philip’s electric toothbrush heads. Before you discard the head, the two magnets on the base can be snapped off with pliers. They are ideal for a host of uses including holding specimens down on to metal bars.

Fig. 5.10 Pupa of a worker wasp; live specimen inside the nest. Both twin macro flash heads were covered in a cone of plastic to diffuse the light so that the translucent pupa would not reflect light. The soft light does not produce a really sharp image but it suits the subject. (×4 magnification, Canon 65mm MPE with diffused macro flashes.)

Fig. 5.11 Use of dessert bowl diffusers over twin macro flash units. A was taken with the diffusers and B without any diffusion, otherwise this small bud of a wild onion was photographed exactly the same. (×3 magnification, Canon 65mm MPE lens, composite of 28-image stack.)

Fig. 5.12 Male doli (short for dolichopodidae) fly showing a close-up of the large genitalia. This small fly has a very metallic exoskeleton and simple diffusion was insufficient to stop reflections and haloes. A long cone diffuser was fitted to the macro flashes and additional paddle diffusers were placed in front of the specimen. (×5 magnification, Canon 65mm MPE lens with diffused twin macro flashes, composite of 68-image stack.)

LED LIGHTS AND CONTINUOUS ILLUMINATION In the four years since the publication of Extreme Close-up Photography and Focus Stacking, LED lighting has notably improved although the IKEA Jansjo lamps discussed in the book remain a reasonable and practical lighting option. They too have improved as some of their original ones, six years on, are now producing a warmer, yellowish light which needs correction in photo editing software. Six years ago they were fine. Whether those bought today (and incidentally still at the same price) will yellow in time is difficult to know, but they now have different LEDs inside. Brighter but more expensive alternatives worth looking at are the super LED lamps

with the Trond brand, found online.

Fig. 5.13 A simple LED control system, top left, for running various LED lights. The black coil is how strips of LEDs can be bought and cut up to line containers such as the one with the illuminated petri dish (thanks to Carel Sartory who made it). The single large LED (bottom right) has been glued to an old computer CPU heat sink. Although LEDs do not get overly hot, some single, powerful LEDs like this one do give off heat and so this can be advisable. Note that the dimmer works as a switch, as dimming can produce a wave effect in the light. LEDs now make up the main form of lighting available. The old tungsten lights

produce a very yellow light and are not worth using because of this and their lower power. Halogen is still quite usable and commonly found in microscope lamps. If you enjoy some DIY, there are some excellent bargains in making your own bespoke system. LEDs have a great longevity and so the bulbs will not have to be replaced for years. Additionally they use little power and can be run from the mains or a battery. For the latter, you can use the same one that is used for driving an auto rail like a StackShot in the field. The easiest option is to buy an LED driver, which reduces the voltage to less than 12 volts. Attach a long cable with a plug for the mains at the other end. Ideally connect the driver up by a short lead to a low voltage socket so that any of your LEDs can be plugged in. You may decide to run this via a low voltage switch or dimmer. Dimmers do not work well with LEDs and if they are set to low power, the light will not be even, but will have a wave-like effect (Fig. 5.13). While a wide range of bright, bare LED bulbs are available for most situations, one great idea is to use flexible LED strips which comes in coiled lengths of 5m. Most are also waterproof as the strip is sealed, making it perfect for lining dishes to illuminate aquatic species. The one illustrated here was made by Carel Sartory and this is just one of many types that he creates. The base unit is just a flat plastic cap painted black inside. The LED strip lines the inside lip with the LEDs facing inward. The wire connections pass through a hole in the side and are sealed in place with silicone sealant. Small dishes placed inside the cap can then be lit and the resulting photograph of translucent aquatic organisms produces an image close to that achieved with dark field illumination (see below). LED lighting will not be as bright as a flash and longer exposures will be necessary. It is for this reason that they may be overlooked and used purely to provide working light to set up everything for photos to be taken with flash. However, there are situations when a ring of LED, either bought or homemade, or even a flash can be ideal for backlighting.

BACKLIGHT, DARK FIELD, ‘STOP’ LIGHTING AND ADDING COLOUR Fig. 5.14 shows the use of polystyrene as a brilliant diffuser by bouncing a flash light off the surface on to the subject. This is particularly useful to backlight a subject and so allow transmitted light through the specimen. The flash is good for freezing action, which can be a problem with aquatic organisms. Overall, specimens needing backlight are those that are translucent or need the edge defining with a rim of light. The dish surrounded by LED strip light will provide a good start, but to achieve good backlighting, a stage made from glass is best. In fact, as we will see, a glass stage is a really useful tool for various lighting. Instead of the flash and polystyrene, there

could be a standard ring light or flash beneath but leaving it like that is missing a trick.

Fig. 5.14 A glass stage for back lighting with a wedge of polystyrene beneath to diffuse the light from the falsh on the right. The green liquid is supermarket window cleaner for keeping the glass spotless.

Fig. 5.15 Dark field lighting. The objective lens visible is attached to a vertically arranged bellows. The specimen in a dish on the glass is surrounded by a black ring, to shield it from stray external light. See Fig. 5.16 for a diagrammatic representation. An LED ring or flash from below creates considerable stray illumination and much of the specimen will be lit with some of this bounced light coming from the side. Dark field illumination is where the specimen is only lit with a ring of light from below. Fig. 4.1 is one example. There are a number of variations on how this can be achieved. The set-up in Fig. 5.15 is part of a DIY ‘system’ discussed in Chapter 6 but here, a piece of glass is held around 7cm above the desk. Under the glass is the ring light sitting on top of black card, so that the middle of the ring is completely black. To help maintain the ring of light, a black tube sits inside the ring; here, a plastic drainpipe cut and painted matte black. On the glass is placed a card with a hole cut in the middle and the specimen placed in the centre. The card here is white, but the area surrounding the cut in the card has been painted black. Then around this cut circle is another short piece of black pipe to complete areas should now have sufficient contrast as they are illuminated.

The camera lens needs to be able to fit easily within this upper black tube and in the illustration, it is an objective on the end of bellows held vertically in place. LED ring lights are widely available and those with a mains connection through a rheostat control are best. Alternatively, one can be made using the flexible strip LEDs. A ring flash will usually work, but some are not complete rings and result in small gaps of light. LED ring flashes are appearing on the market at low prices but expect the very cheap ones to have limited power and the flash duration to be quite slow. This means they will not freeze any action like a true flashgun. With a little ingenuity and careful use of polystyrene for diffusion, a ring can be made based on Fig. 5.14 but using two flashguns at opposite ends.

Fig. 5.16 Diagram of the dark field set up in Fig. 5.15. Achieving dark field on a microscope is quite straightforward and has been well documented. It is based on a ‘stop’ filter placed in the path of light before entering the condenser. Some universal condensers have these permanently fitted; they are just

rotated into place instead of bright field lighting. If not, filters can be bought or simply made. As a ring of light is required, the filter has a large black spot (the stop) in the middle surrounded by a clear area that produces the light ring. There should be the totally blacked-out arrangement, so that the specimen is only lit from below. The specimen will then have a dark background and the transparent a filter or stop holder that swings in and out under the condenser. As the transmitted light passes up from the field diaphragm, the rays will be blocked in the centre and only a cylinder of light now enters the condenser to be focused on the specimen. If a blue spot is used instead of the black in the centre, a particularly super effect can be created, referred to as Rheinberg illumination.

Fig. 5.17 Examples of homemade stops for microscope sub-stage condensers. Thanks to Carel Sartory for making these ones by printing a template on to acetate sheet. These are also used in the ‘stop box’ as well. This stop holder can support any number of variations on this theme. Replace the black stop and clear rim with colours like a green stop and yellow edge. This produces a green background and a yellow light to the specimen. A red stop with blue rim yields red background and a blue specimen. The size of the stop is important and

a complete guide for making Rheinberg discs can be found on the Quekett website (www.quekett.org/resources/ rheinberg) along with some excellent photographs to illustrate the different affects. All of these techniques, whether for microscopes or other extreme close-up methods, are to improve the contrast in transparent or translucent specimens. When using these centre stops on microscopes, Köhler illumination still needs to be set up first. The variations mentioned here are purely ways of adjusting the Köhler light to improve the ability to see the specimen more clearly. Microscopists have tried for centuries to find ways of doing this and continue to do so now. One simple way is just to create a deliberate uneven lighting (oblique lighting) which introduces a shadow and so forms a three-dimensional image. This can be done by putting a small piece of black tape on the stop holder under the condenser to blank a quarter of the light passing upwards to the specimen. Alternatively, cut a circle of clear acetate to fit the holder and then blacken (using tape or card) a small wedge in the circle like a slice of pizza. Try different sizes of wedge, but ultimately you are forming an uneven light that shows up transparent material within the specimen to give a 3D image. All of these techniques can be modified for use.

STOP BOX Stop filters should not be limited to microscopes but used as a general lighting technique. There are numerous ways of setting this up. My example in Figs 5.18 and 8.15 show a homemade ‘stop box’ made out of wood. On the box base is fitted a bright LED and the only outlet for the light is a hole above it. On the hole sits the stop filter, held in place by a large convex lens (which came out of an old enlarger). This directs the light through a hole in a stage fitted approximately 7cm above. On that stage sits the specimen. The lens needs to be moved around a little until the effect is at its best. The specimen is viewed from above. If a different filter is needed, the lens is lifted off and a new one placed underneath.

Fig. 5.18 Stop box. A: powerful LED light is focused by the large lens (from an enlarger) on to a specimen. A stop filter is placed under the lens; B: rove beetle, squash preparation after maceration. The transparent subject was photographed with transmitted light from an LED stop box where a blue stop with a red rim was used. (HDMI inspection microscope ×20.) From here on, the lighting methods can become increasingly more complex and while they have been designed for microscopes, all can easily be crossed over and utilized with our extreme close-up techniques.

Cross polarization is a method where two separate polarizing filters are used in the light path. When using photographic filters, ensure that they are the same type, for example, circular polarizers. One is placed beneath the condenser or light source and so polarizes the light entering the specimen. The second is ideally placed above the objective lens and is referred to as the analyzer filter. This one stays fixed in place while the one below the specimen is the filter, which is rotated. If you hold two filters together against the light and rotate them, the light dims until it goes black. Keep rotating and the light reappears. A single polarizer is traditionally used on a camera in bright sunshine to minimize polarized light so that clouds stand out and the sky is darker. It helps to reduce reflections on vegetation.

Fig. 5.19 Cross polarizing light to create colour in images of minerals. A: cinchonine, a substance extracted from the bark of a Peruvian tree and used as an anti-malarial drug; B: vitamin C melted slowly on a hot plate and allowed to cool. (Both ×40 with a Sony A6500.) Some biological material but mainly minerals are optically anisotropic. This means that when hit by polarized light, the plane of the polarized light is rotated as well as altering the nature of the light. In our set-up, by sending polarized light into the transparent or white specimen if it is anisotropic, the light is twisted and altered. The

analyzer filter determines what light will eventually be passed on to the camera. Results are difficult to predict and this is done purely so that the specimen stands out with colour and contrast, where before there was none. The samples shown here are easily obtained minerals, which can be made up into supersaturated solutions and melted on a hotplate. When allowed to evaporate or cool from the hotplate, the crystals grow quickly and it is a case of tracking them under ×20 to ×80 magnification and rotating the lower polarizer to create the colours. As you rotate, so the colours change through greys to possibly every colour of the rainbow and then back again. Areas outside the crystals might be just black and to boost colour in those parts, you can alter the light above the first polarizer with a thin layer of polycarbonate. The easiest example of that is the clear lid of a CD case. Moving and twisting that will give every colour from pink through purple to green and yellow. Remember, this is just to produce contrast and colour, but the results can be so addictive you just want to keep making crystals. Although an easy technique, the problem most likely encountered is where to place the analyzer filter. Most research microscopes and even some student ones have a slot in the tube between objectives and eyepiece where one can be inserted. If not, it needs to go above the eyepiece and before the camera sensor. For bellows fitted with an objective it can be possible, with Blu Tack or some other temporary adhesive, to try mounting a small-diameter filter within the bellows at the end nearest the camera mount. Especially popular in the 1970s and 1980s were mirror lenses for SLR cameras. With very wide barrels to hold the mirrors, they have special filters to fit in the base rather than screw into the front. This means they are small, with around a 30mm diameter. A set of filters of all colours and a polarizer could be bought. They are still available second hand on auction sites and are ideal. Spike Walker is an award-winning British photomicrographer who has extended the Rheinberg illumination by introducing different colours through varying techniques (referred to as Spikeberg). A particular hallmark of his work is extreme close-ups of crystals. He takes supersaturated solutions of substances like vitamin C and, as they crystallize, cuts lines into them so that he creates incredible diversity of shape and pattern. Spike then uses cross-polarization as part of his Spikeberg illumination to produce intense colouration of what would otherwise be quite uninspiring material. Check out some of the galleries available online of his stunning collection of photographs. As the magnification of a specimen increases, so lighting a subject from above can be difficult. A ×10 objective on extended bellows can have a working distance of around 4mm. Much smaller and lighting can become a major issue. On some microscopes it is possible to set up what is called epi-lighting using specialist equipment. In brief, light comes from a source at the back and passes along a horizontal tube above the stage. This connects at right angles to the vertical

microscope tube above the objectives. The specialist objectives are much fatter than other objectives, as they have an outer sleeve with a gap between the inner objective and the outer sleeve. The light passes down this gap and at the bottom is reflected on to the specimen by a mirrored surface. In this way light is delivered straight on to the material. Filters placed in the path of the light can be added to create dark field and other modifications. For microscopes, this is an excellent way of illuminating opaque and dense subjects and is important in mineralogy. The principle of epi-lighting is relatively easy, if rather fiddly to get right, with bellows and objectives. Alfred Blaker (1977) includes epilighting in his detailed descriptions of ways of adapting microscope illumination methods to macroscopic and extreme close-up techniques. He refers to this as ‘beam-splitting of axial light’. In its simplest form and the one used here, Fig. 5.23, black sticky tape (Gorilla gaffer tape works well) is rolled into a tube and cut at approximately 45 degrees so that a thin piece of glass such as a microscope slide or coverslip can be stuck in place. The tube is the diameter of the lens to be used (such as a ×10 objective) and stuck on to the end of the lens. The photograph is taken through the glass as it is focused on the specimen. A parallel beam of light is shone on to the glass from the side and some of the light reflects down on to the subject, brightening up any dark crevices where the side incident light cannot reach. It is important that the inside of the tape is black so that there is no internal reflection. With care, this simple arrangement can provide good epilighting on an ad hoc basis. If you want to develop other DIY methods of lighting and more permanent epiillumination, Blaker’s book (although out of print and based on film and outdated cameras) is well worth a look if you can find a copy.

Fig. 5.20 Epi-lighting illuminating the terrestrial alga Phycopeltisanundinacea, on an ivy leaf. (×80, photo taken by Mike Crutchley.)

Fig. 5.21 Epi-lighting set-up on a Zeiss microscope. Light passes from a source on the right and enters the main tube at a right angle through a prism above the specialist epiobjective.

Fig. 5.22 Epi-lighting to photograph a grub in a mud and silk case (unknown species). A number of these tiny mud globules were collected and upon cutting one open, they were found to have a grub inside. Epi-lighting was achieved by using the set-up shown in Fig. 5.23 and this illuminated the shadow areas in the lower part of the mud case. (×15 with bellows and ×10 objective.)

Fig. 5.23 Diagram of a homemade epi-lighting method (after Blaker 1977).

FLUORESCENT LIGHTING If you have had your banknotes checked in a shop, you will have seen fluorescence in action. If you have not tackled this form of photography before, you are in for a whole

new realm of image specialism, which can merely be touched upon here. The subject is not confined to extreme close-up work but can be used for virtually anything from fashion through to nature. The principle is that under certain wavelengths of light, much that could not be seen appears bright and ghostly, and sometimes in amazingly weird form. This is typical of ultraviolet light when it strikes certain materials, called fluorophores, as it causes them to become excited (the correct term) and fluoresce. It is this fluorescence that can be photographed. One of the most spectacular sights is to photograph a scorpion in UV light; in fact, it is easiest to find them in the first place with a UV torch at night. Quite a few arachnids reflect UV, as well as other nocturnal creatures such as millipedes and some snails. Many marine organisms demonstrate fluorescence. In warm seas like the Mediterranean, swimming at night can cause the water to suddenly produce a cool light, but this is a different process called bioluminescence. While fluorescent light can create dramatic photographs, this form of illumination is again about showing up features that would otherwise not be visible.

Fig. 5.24 Eyes of a Spanish tarantula. The specimen was collected in southern Spain and preserved thirty-five years ago. They are small spiders with a body almost 2cm long. All are composites of a 25-image stack taken with a Canon 65mm MPE lens at ƒ5.6 (×6 magnification).A: light was a twin macro flash at ISO 200; B: light was a 51 LED torch emitting 395nm light, an eighth of a second ISO 1600; C: light was a single LED Windfire torch emitting 365nm light one thirtieth of a second ISO 6400, and note the clean image with no general violet found in the 395nm light. Yellow protective glasses were worn during the use of the torches.

The visible spectrum to which our eyes are sensitive is in the wavelengths between 400 and 800 nanometres. The latter is the red end of the spectrum which runs into infrared and on to radio and TV wavelengths. The other end is blue light moving towards X-rays and sandwiched between them is ultraviolet; 400nm is visible to us as violet and below this, light becomes difficult to see as it enters the wavelength of ultraviolet. A large number of UV torches are available for the price of a cup of coffee. They are typically used for checking banknotes and stains on carpets, but will show up arachnids and other natural products in the dark in a rather violet glow. They are inexpensive as they are not especially bright and because they are only just into the UV spectrum at around 395nm. Brighter ones with 50 or 100 LEDs are best but ‘bleed’ into the visible spectrum and will not provide the cleanest of images. This can be improved with filters to remove the visible colour. The ideal LED light for nature photography is around 365nm, well beyond the visible spectrum, so that a clearer florescence is produced with cleaner, more vivid colour and no violet glow. Expect these to be around twice the cost or more. The Convoy S2 range is good but a bit pricey and at half that cost is the Windfire torch, which is more than adequate for the job. As 365nm is not in the visible spectrum, the torches are sometimes labelled as producing ‘black light’ because all that is emitted is a vague glow. Having two torches, one of each wavelength at 365 and 395nm, can be useful. The latter is sufficiently bright to help find material at night, while the other can be used for photography. If you start working with ultraviolet, be aware that UV is dangerous to your eyes and caution should be exercised not to look directly at the light. Snow goggles and driving glasses should offer UV protection if you have them and if not, buy a pair of protective glasses to be worn while you are working with the light. Some of the better, more powerful torches come with the yellow glasses. The 365nm torches can be used just like the LED lamps discussed earlier in the chapter, although you will need to work in a dark room so other light does not affect the imaging. The problem is the amount of light the torch emits, which will require the use of long exposures. It is preferable not to expand the ISO too much (although this is often necessary) as the dark areas will produce unwanted noise in the digital image. RAW files will be better than using JPG so that in processing, noise can be reduced. The beam of light can be quite narrow which means that you might find yourself ‘painting’ the light on to the specimen during the long exposure.

Fig. 5.25 Autofluorescence. Many organisms, especially marine creatures, show fluorescence when struck by certain wavelengths of light; 450 nanometres of blue light cause the red and green to fluoresce, bringing out the detail. A: hydroid Obelia and B: a colonial hydroid covered in diatoms that show up red. (Both photos taken by Mike Crutchley.) Buying individual bare LED bulbs that emit specific wavelengths enables you to enter an even wider world for both microscopes and macroscope situations. For the latter, the bulbs can be attached to the inside of a white box with a hole in the top on which to place the specimen. Any of the previous set-ups like the stop box could be

adapted to contain the UV light. Depending on the emitted wavelength, it can help to minimize any stray visible light by using a yellow filter. Any photographic yellow filter will do for this, whether an expensive brand or a cheap square from an eBay shop. This ‘analyzer’ filter allows just the excited colours through and instead of a general glow, the image will be dark, showing only the specific proteins that have fluoresced in red or green. The Obelia image in Fig. 5.25 has been photographed twice; once with normal light and once with blue light. Then the two images were merged on the computer. This allows the colours to be seen with the whole creature. Hydroids are an interesting group of primitive marine organisms. When their fluorescence was investigated, it led to several discoveries now used in genetic engineering where the excited proteins are used as markers in the process. Obelia is very common on rocky shores. It is transparent but when a wavelength of light of 450nm (blue) is shone on to it and photographed, green proteins become ‘excited’ by the light and show up as green. This is most likely to contain DNA. Chlorophyll under 400–450nm is excited to become red and when out looking for scorpions with the UV torch, young plants and some tree lichens can stand out in deep red. Fig. 5.25B has diatoms all over the surface in red which otherwise would not be visible. True fluorescent microscopy goes way beyond this simple level of photography. Traditionally the light source was a mercury vapour lamp requiring careful operating conditions, including the slow warming up and shutting down of the light. The light source emits a variable range of frequencies including visible blue light. To use this light, the wavelength is adjusted with a series of different filters. Modified microscopes like the Leitz Ortholux have the light arriving (like the epi-lighting) into the microscope tube via a specialized analyzing section containing filter cubes. To increase the chance of specific proteins fluorescing, chemicals can be added to materials. These fluorochromes, such as acridine orange, react with particular proteins and make them fluoresce. Just a tiny amount of acridine orange will make the nucleus stand out orange in cheek cells scraped off from the lining of the mouth. It sounds painful, but it is very simple as these cells are flaking off all the time and all that is required is the use of a spoon handle to rub up and down the inside of the cheek. The pale drop on the end of the handle is then placed on a slide with less than a millilitre of acridine orange, causing the nuclei to slowly appear in the cell under a ×10 objective lens. If you want to investigate further into nature photography using light beyond our visible spectrum, look at the work of the photographer Adrian Davies. Lighting is an exciting way to extend your view of extreme close-up photography and you may need to check on the references at the end of the book. Throughout this chapter we refer to peripheral equipment to support lighting and this needs more explanation.

Fig. 5.26 Proboscis of the house fly, removed and placed on a glass slide in glycerol. (×35 magnification. Photographed with a Canon 7D mk2, bellows and ×10 Vickers objective, dark field illumination and with a fill-in flash from the side.)

Chapter 6

Support and Preparation

ow we seem to have all the elements necessary to create our image: camera and lenses with suitable light. Except that with extreme close-ups, those are just the fundamental, expensive parts. On a practical level there is a whole host of small but essential items left, such as anchoring the camera to a solid support to stop vibration and blur. More complicated can be supporting the specimen and if we are going to magnify the subject, how do we manage to see what to photograph? We also need to consider whether we are photographing outdoors in the field or in an indoor studio.

N

Fig. 6.1 From marine plankton, a leaf worm larva on a gelatinous colony of the diatom Phaeocystis. Photographed in a dish of seawater with magnesium sulphate added to slow the worm down. (×10 magnification Canon 7D mk2 with bellows and ×4 objective, dark field.)

SUPPORTING THE CAMERA

In the field a tripod is the default support but not all tripods are the same and for the better ones, the head has to be bought separately. Carbon fibre will be more lightweight than aluminium, although more expensive. The chances are that you will be carrying a fair amount of equipment and possibly the difference in weight is not worth the extra cost. Having a tripod that has some ability to shoot at lower angles is really useful, either by virtue of the centre column being held horizontal or the legs collapsing out flat. The old style Benbo is brilliant for that, if a little heavy. The new Benro GoPlus series is almost as good. Alternatively there are plenty of excellent tripods from the folds of Manfrotto, Gitzo and Vanguard.

Fig. 6.2 Manfrotto three-way geared head for a tripod. Far more important is the head that you buy for the tripod. Ball and socket types are popular and simple to use. However, they have one major flaw unless you buy a very expensive one. As you set up the rail, camera, lens and possible lighting, all on the head, the weight can make it a little difficult to manoeuvre into position. Then when you find the rough focus point, you lock off the head so that it does not move. Having locked it, you let go of the camera, only to see the play in the system causing the set-up to drop, only a few millimetres but sufficient that it is no longer in the

correct position.

Fig. 6.3 WeMacro vertical stand. A: the bare stand; B: fitted with a StackShot auto rail with camera. Note the addition of two Zeadio 11in magic arms for holding flash heads. Having tried numerous types of tripod head, the most recommended one is the three-way geared version. They take a little getting used to but the best ones have a quick release for each of the three planes. By pulling a lever, the head moves the camera in that plane and so the approximate position is established. Then a knob within the lever is rotated for fine control so that the head and camera move smoothly to the exact position required. All three knobs, one for each plane, will require finetuning but after a ball-head, the ease of setting up a shot is fantastic and you will never want to go back.

Arranging a tripod for close-up work is the one activity most likely to make people irritable. Handholding a camera is virtually impossible as you pass from macro into extreme close-ups. Unfortunately, when working in the field there is not much in the way of options. Bringing your tripod indoors will take you beyond irritable as it becomes even more difficult to set up against desks and tables. It is not impossible and for occasional work you might be able to get accustomed to how to do it. If you have sufficient table space, the best decision is to make a small studio with an optical bench. In its most simplistic form this is a solid construction, maybe made from small planks of wood and metal on which to bolt the focusing rail. The camera moves along this bench horizontally back and forth towards the specimen, which is mounted on the same plane. It does not have to look good, just be solid enough to take and support the camera equipment. This is all right if the specimen can be held out in front of the camera but there are plenty of instances, for example with aquatic creatures, when it is more suited if the camera is held vertically to run down to the specimen held near the base. While a DIY option is possible, vertical benches are not so straightforward to create in a solid enough construction. Both enlarger and copy stands can be found on the second hand market. These have a solid base with a vertical column. In the case of the enlarger, that can be removed to be replaced by the camera, which is cranked up and down. The copy stand is slightly easier as it will definitely have the fitting for a camera and can be more substantial. If it is an enlarger base, buy one that is large and solid from a well-known make. Copy stands from Kaiser are good and sell on eBay for around £150. The best buy is a new stand by WeMacro, a company that produces the Chinese auto rail. For around half the price of a second hand Kaiser copy stand, the vertical stand is ideal and does not take up a large space on the desk. It is very solid and has a variety of threaded holes to take accessories like magic arms, which can be bought on all auction sites. Without too much effort the column can be removed and bolted flat for horizontal shooting.

Fig. 6.4 Homemade optical bench based on a Prior microscope stand so that it can be in either vertical (A) or horizontal (B)plane. The stand is bolted securely to a decking plank of wood at the end so that in the vertical plane, suitable lighting can be placed underneath. A WeMacro auto rail with camera is attached. The strange feet are exwindow frame plastic, which is rubberized and provides stability and grip to the surface. My alternative to all of these is an optical bench based on a flexible microscope stand. The one in Fig. 6.4 is the stand from an old school Prior microscope that can be picked up second hand quite cheaply. The rack and microscope tube are removed and replaced by a short length of steel. This has been predrilled to take a bolt that will enable the attachment of an auto rail. The feet of the stand are drilled so that the base can be firmly bolted to a solid and stable bench. Here, a length of wood decking, around 80cm long, forms that base which is itself fitted with rubber and plastic feet so that it does not move. The microscope stand has been fixed right at the end of the wood so that when it lies horizontal, the long base gives stability. Moved to the vertical, the base still gives sufficient weight at the other end so that it

does not tip. Also by being at the end of the wood when the microscope is in the vertical position, the camera lens is pointed down to an open space for placing a stage for dishes or different lighting, like dark field.

MAKING A STAGE A camera on a rail can go back and forth for focusing but there are no adjustments in the other two planes. With the camera equipment stabilized on a stand in the studio, next is setting up a stage on which to arrange the specimens. Vertical or horizontal arrangements depend on the subject matter and this may alter the stage arrangement. There is nothing complex about the stage; it is a homemade job with many possible variants. The basic stage for a horizontal camera position will have a flat area that can move up and down, so the specimen can be carefully positioned. Lab jacks are available in different sizes and these, with the turn of a knob, will do just that. The likelihood is that there will be insufficient space at the top and this will need to be extended by bolting a larger stage area at the top of the jack. UPVC from roofing soffits works well and makes a wipe-clean surface. This can be drilled to make holes in which lighting and other accessories can be fitted. Blu Tack and other temporary adhesives can be used to attach things to a UPVC surface as well. This may be necessary to prevent the movement of dishes and small tanks. UPVC is also easy to clean and if you work with aquatic material, to wipe dry.

Fig. 6.5 A stage for use mainly where the camera is held horizontal in a studio. Made from a piece of UPVC, it is bolted to the top of a lab jack (here, raised high for the photograph). Holes have been drilled to take the blue pipes (flexible plastic water coolant pipe hoses), which have wires running through from bright LED bulbs down to a connector and switch (bottom right). The black clamps on the edge and the two suckered clamps on the stage are for holding the macro flash heads. Blu Tack is always to hand on the stage and a spare, smaller lab jack is shown bottom left, used on travels. Holders for specimens and diffusers can be placed on this stage with the smooth surface making lateral movements easy. While specimens can be placed on stones, twigs and other natural materials (keep a small collection handy), they may need to be on more flexible holders. ‘Helping hands’ can be bought in bargain shops and online. They consist of crocodile clips mounted on flexible joints, attached to small metal bars that rotate on a heavy stand. Several sets of these are worth having,

although homemade ones with a crocodile clip fixed into a wooden ball work well. The wooden ball sits snugly in a suitably sized lid from a small jar. This allows the ball to rotate and move about.

Fig. 6.6 Helping hands for holding specimens, background cards or wire paddle diffusers.

Fig. 6.7 Reverse forceps. The basic forceps with the blue rubber grips have to be held in a helping hands clip, but the best ones available are the larger and substantial form with a heavy base. The lock joint in the middle allows precision movement and control. Other forceps, both coarse and fine are useful along with a fine brush. One of the most useful devices is a set of reverse forceps, mounted by a lockable joint to a heavy base. The one shown in Fig. 6.7 is readily available online and has a strong action. By squeezing the sides the ends open to grab the specimen. Upon releasing the pressure they grip and hold the specimen very tightly. The joint enables good lateral rotation as well as up and down. A combination of this with the lab jack gives excellent positioning. With a total length to the middle of the reverse forceps base of 20cm, you will need a fair amount of space on the stage to accommodate this along with possible lights and diffusers. A stage made from UPVC is good for attaching clamps for lights. While tripod makers produce flash clamps, they can be expensive. By checking auction sites like eBay, you can find a great diversity of very cheap ones but they will not necessarily have ‘flash’ or even ‘clamp’ in the title. In Fig. 6.5 there are two different holders suitable for supporting the twin macro flash. These are sold for sat navs or phones. The clamps (sold for bicycle handlebars) fix to the sides of the stage while the others have a large sucker on the bottom for fixing to windscreens, but will attach just as well to UPVC.

The stage for a vertical stand could just be a variation of the horizontal one. As there is already a base present, such as the one seen in the WeMacro stand, some accessories could attach directly to that. For example, magic arms can be attached. The Manfrotto version is more than five times the cost of those like the Neewer or Zeadio 11in arms. One end screws into the base and the other end holds anything from flash to diffusers or monitors. They are completely flexible and can be moved into any position, until the large knob in the middle is tightened and the entire structure locks up. By setting these towards the back, a simple flat stage is placed on the front of the stand. For vertical stands an X–Y axis mechanical stage like those used on microscopes can be fitted, although the right type needs to be sourced, as some have the control knobs located below and space is at a premium. The WeMacro stand comes with an X–Y axis arrangement, which is essentially a pair of small manual rails, one attached to the other. A crocodile clip is fixed to the middle. Alternatively the reverse forceps will work just as well on this stage, as will a small petri dish. Like any stage, you will need to think of the background. Keep a good collection of coloured cards of different sizes, which can be held behind the subjects using the ‘helping hands’. In most cases, these do not need buying as plenty of material arrives in the post or with shopping that is normally discarded. With extreme close-up photography, the size required is minimal. Neewer equipment tends to arrive in good quality black boxes producing excellent card squares of 6 × 4cm or 10 × 8cm. To place under petri dishes on the stage of a vertical mounted camera, coloured plastic (blue is most useful) is preferable as water spillages from the dish can be mopped up. A roll of jumbo paper towel is always kept handy. Black, blue and green are good colours to use for backgrounds but do try complementary colours, even within the same backdrop. One way is to take white card and by using watercolour paint, produce a series of washes with a combination of colour. Remember that in most cases these cards do not need to be large and it is a good idea to produce different sizes so they are at the ready. In addition to all of these items, a workbox with divisions for keeping materials separate, obtainable from supermarkets, is great for storing all the small and loose things. Tools like fine forceps and scissors. The most useful scissors are a very fine type that are controlled by squeezing in the middle to make them cut. Mount needles into a dowel of wood by drilling a very small hole in one end and gluing the blunt end inside. These are good for moving tiny specimens around. Small boxes of different sized pins; a reel of fine cotton thread; pipettes; thin wire; Blu Tack and a collection of small but powerful magnets. This last one may sound odd but they are excellent for quickly holding down leaves and fabric backgrounds to the arms of ‘helping hands’ or other metallic surfaces. If you have access to old Philips Sonic toothbrush heads they have two very strong magnets, which can be removed with a pair of pliers.

Fig. 6.8 Dusty eye of a horsefly. Insects are invariably covered in dust, pollen and debris. The white detritus covers the eye and some is reflecting the flash, causing the exposure to burn out the detail. (×12 magnification. Canon 7D mk2, bellows with ×4 objective, twin macro flash, composite image.)

DEAD OR ALIVE?

One of the most important accessories you can have is a hand lens. The starting point for any photograph will invariably involve looking carefully at the specimen to check exactly what it is you are homing in on to photograph because almost certainly it will not be visible to the naked eye. Sometimes at this point you realize it is damaged or very dirty. Insects and the like will pick up an incredible amount of debris, which is why when watching a living fly you will see they spend a good deal of time cleaning themselves. If you are working with dead material it is possible to clean it with a small ultrasound cleaner, the type used for cleaning jewellery. A fine paintbrush with the tiniest amount of solvent can be combed through the insect to remove some debris. It may be that one side of the creature is better than the other and once the best viewpoint is determined, the subject can be set up. At some stage you might find a hand lens is just not good enough for this and that is where a low-power stereomicroscope or an HDMI inspection microscope comes into its own. These will be easier to set up small specimens in a holder and then manipulated into the right angle for photographing. Do you use live or dead material? In the previous book Extreme Close-up Photography there was an emphasis on looking after the live organisms and photographing them whilst alive. The majority of websites on this topic discuss dead specimens and how it is impossible to focus stack live creatures, especially insects. This is incorrect, as it is perfectly possible to do large stacks of extreme close-ups of many species of animals. There are certainly plenty of difficult or impossible species, but not all of them are. As usual, the answer is that it depends on other factors. This book is about having the ability to photograph any organism needing magnification between ×1 and ×100 whether from land, freshwater or sea. The higher the magnification, the greater the chance that the animal will have to be killed, or ‘euthanized’ as some people prefer to say. Depending on the circumstance, try not to automatically assume that a specimen has to be killed. The hawk moth in the next chapter was magnified ×5 and one scenario looks at the process of focus stacking one of its eyes. Throughout the process, the moth was alive and released afterwards in a healthy condition. In so many circumstances it is clear to a naturalist when a creature was photographed dead or alive, the eye being one of the first places to show this. A dragonfly’s eyes show incredible colour and detail when photographed alive, but this will start to diminish the moment it is killed. Fig. 6.9 shows an extreme close-up of the eye and head of a prawn where the square optic nerve is clearly visible. That will be the first thing to go on death and so for marine material as well as terrestrial beings, keeping them alive is an important consideration. Many people that say killing insects and others is the only way forward to photograph them is often due to laziness as it can take time and patience but it is very rewarding when it works.

Fig. 6.9 Head of a live prawn, Palaeomon serratus. If the animal were dead, most of the detail in the eye and the star-cells (chromatophores) would quickly disappear. Photographed in an observation cell of seawater backlit with a flash and a fill-in flash at the front. Various card backgrounds were tried but the black produced the contrast necessary to see the rostrum between the eyes. (×4 Canon 7D mk2 with 65mm MPE.) Before killing the animal for extreme close-ups, learn about the biology as it may well be a species that under certain conditions will sit still and pose. Second, check that it is not a rarity and that there are plenty of others around. Please think of conservation and limit the number of creatures that you take. Aquatic life can nearly always be photographed without killing the specimen below around ×8 magnification. Above ×10, most creatures become difficult to photograph alive. Ultimately, it is down to your conscience and care.

HOLDING CONTAINERS AND OBSERVATION CELLS

If the specimen does need to be euthanized, the best method is to place it in the freezer while contained in a tube. Ten minutes can be sufficient time to subdue an insect, bring it out to photograph and eventually release it alive. Wasps have been kept up to twenty-five minutes in a freezer and still they come back to life. Soon after removal from the freezer, specimens develop condensation on shiny surfaces, especially the eyes. Obviously prolonged exposure to freezing conditions will eventually kill the animal. If a freezer is not available, the usual method of quickly dispatching an insect or similar invertebrate is to use a few drops of ethyl acetate on cotton wool or a tissue and place it in the tube or jar with the animal. This kills them quickly and keeps them relaxed to then set them up for photography. Aquatic life should be treated differently but some of what follows may be suitable for some terrestrial live specimens as it is important to make up suitable containers, holders or slides. We mentioned above that petri dishes were good for presenting specimens to view from above. If you are in IKEA buying the Jansjo LEDs, pick up a stack of T-light glass dishes. Normally twelve are stacked one on top of the other in a packet and these are excellent value 4cm diameter dishes for holding aquatic material. Add a minimum of water to just cover the specimen.

Fig. 6.10 Foreleg of a male great diving beetle, showing the sucker for holding the female. Taken live in an observation cell, there are white protist animals on the leg. Note the detritus in the water. (×3 magnification.) For really small material, instead of placing it in a dish, use a microscope glass slide, especially one with a cavity in it. Cavity slides may not be deep enough and so a ring of Vaseline or petroleum jelly can be made around it. The water is added with the specimen and a coverslip placed on top to produce a flat surface, otherwise a bowed meniscus can result in distortion. You can make your own wells on a slide by gluing washers or plastic rings to the glass. These need only be a centimetre or so across in diameter and a variety of different thickness are useful to set up. The idea is to choose a suitably sized space that holds the specimen easily without damage, but which creates some limit to its movement. Some aquatic species such as crustaceans will move whatever the space, whether it is legs or antennae. There are ways of stopping this by a physical ‘restraint’, an increase in the viscosity of the water or a chemical that will narcotize them.

Fig. 6.11 Planktonic larva of a sand mason worm, Lanice. A live specimen photographed on a slide with a coverslip, which was held up by four Vaseline blobs at the corners. Gentle pressure trapped the worm tube so that it could not move. (×14 magnification. Illumination with a flash beneath and one above. Canon 7D mk2 bellows with ×4 objective and twin macro flash.) One technique shown to me by Wim van Egmond (check out his incredible images and video at www.micropolitan.org) was to place four spots of Vaseline on a slide where the four corners of a square coverslip will go. Place the specimen in the middle with water, add the coverslip and then press down gently while looking at the specimen. By pressing carefully with a needle from one side, the specimen eventually becomes trapped by the coverslip, acting like a wedge. Often this is all that is needed (see Fig. 6.11). There is a device, difficult to obtain now, which does a similar thing to the four Vaseline spots. In essence it has a knob that you rotate to pressure down and control the squashing process. Adding a little cotton wool or even tissue will prevent a creature from swimming. Long narrow fibres entrap a creature such as a water flea and keep it in the field of view. Although it does work, the problem is that the fibres often cross over something you wish to photograph. When dissolved in water, methylcellulose produces an increase in viscosity as it becomes like a gel so the animal struggles to swim. Methylcellulose is easiest to buy in the form of wallpaper paste. Once made up, the paste needs to be kept well sealed in a container with a minimum of air, as over time it will lose its viscosity. Better still keep a small, sealed bag of the dry wallpaper paste granules handy in your tool box and make up as necessary. Ideally source the type without fungicide, although it does not seem to adversely affect the creatures. If it is a seashore creature, make sure seawater is used. Put the made-up paste in the container and add the animal. This will probably bring in a small quantity of water with the specimen so use a needle to gently move the paste around. Wallpaper paste slows down the

movement considerably and most insect larvae will stop altogether. The paste has no effect on the aquatic creature, as it is more likely to run out of oxygen before that point is reached. A quarter to half an hour is more than enough time before they should be released. Even water mites (which are notoriously difficult) tend to slow right down but they may not stop moving body parts completely. Glycerol or vegetable glycerine is readily available from pharmacists or online and can be mixed with water to increase viscosity. It is not so useful for marine species but can help for some pond life, although it can be sticky and messy to use.

Fig. 6.12 Tentacle of live Portuguese Man o’ War, Physalia physalis. Washed up on the beach, a tentacle was cut off and photographed in a small observation cell of wallpaper paste made up with seawater. Even an hour later the tentacle was writhing but the paste prevented movement long enough for a seven-image stack. (Composite ×6, Canon 7D mk2 with 65mm MPE with twin macro flash, one flash top and one side.) If the animal, like the water mites, is one that will not stop moving you might have to think about narcotizing them. An anaesthetic throat spray may also work. Epsom salts (magnesium sulphate) can be made up into a super concentrated solution and then

drops added to the water containing the animal. Wait for it to take effect as it narcotizes the creature – experience will tell you how much to add. Too much too quickly will kill the specimen, but the right amount will arrest movement and after washing with clean water, the creature should swim off. Other agents like magnesium chloride or alcohol dropped in the water also work. For marine life, few things beat magnesium sulphate and it has been used for years to relax specimens to make photography easier. Do not be in a rush to use these measures of narcotization or restraint, as quite a few species will initially be manic and then settle down. In the References section, there are various sources to find out more about narcotizing agents. As well as Vaseline and wallpaper paste, silicone sealant is useful to make wells and supports for coverslips. Support means just that: a small blob of either Vaseline or silicone in four corners on a slide can be used to support a coverslip so that it does not crush the beast in the liquid beneath. The former is very temporary whilst silicone is a more permanent set-up on the slide. It can be used to make rings as well, although the most useful part of silicone is its use in sticking glass together to make observation cells. The glass in these cells needs to be thin and optically good. To achieve side-on shots you need to set up a minute version of an aquarium. The problem with any tank or aquarium is that upon setting up your camera on one side, the creature inevitably moves to the other, away from the camera. So the principle of these observation cells is that they are wide at the front and very narrow on the side. The simple version is made from two microscope slides sandwiched together, squashing a U-shaped length of Blu Tack. The width of the interior can be varied by squashing tighter or by using more Blu Tack. These are temporary and after a while they begin to leak. This can be improved by using a clip at both ends for the two slides to be held together. An alternative to the Blu Tack is a short length of narrow plastic tubing that is bent into a U-shape and the slides clipped.

Fig. 6.13 Spirobid worm attached to seaweed. The tiny worm projects tentacles from the tube for feeding but movement is too quick for stacking. Magnesium sulphate was added to the water, which relaxed the animal and slowed the tentacles sufficiently for a siximage stack. A small amount of sand or debris can be added before the specimen so that it sinks to the bottom and covers the Blu Tack or tube. A better and more permanent arrangement is to use silicone to glue the slides together. This time you need as many as five slides. The slide that will be the front has the bottom edge glued to a slide laid flat. The slide to form the back of the cell also has the bottom edge glued to the base slide, butting up against the bottom of the front one but the top edge leaning backwards a little to produce a V-shaped section. To complete the observation cell the ends need enclosing, which is where the remaining two slides are used. These will be too long and so if one can be broken in half, the two pieces could then be used instead. The long narrow slides work quite well but it is useful to have different sizes and shapes. The majority of microscope slides are 2 × 7.5cm but it is possible to obtain 4 × 7.5cm, which work well.

Fig. 6.14 Observation cells. Made from old photographic plates and glued together with silicone sealant, a wide variety of sizes and shapes will cater for all situations.

Fig. 6.15 Live water beetle larva photographed in an observation cell. The animal moved and the composite needed retouching. (×4 composite from an eight-image stack.)

Fig. 6.16 A single Bowerbankia animal feeding. Commonly known as sea mats, they can cover areas of seaweed and feed on detritus in the water. They open like this very quickly and then shut down, so stacking is almost impossible. (In a small observation cell, ×12 magnification, Canon 7D mk2 with bellows and ×4 objective lens. Flash.) For quality and thinness, an excellent alternative is to use old photographic plates. They can be obtained as 4 × 4cm or approximately 10 × 8cm. Both produce very good observation cells and being photographic plates, the optical quality is perfect for this type of work. You could try to find a small collection of these plates on eBay, either unused or with a developed negative present. To remove the emulsion, place a plate in a washing up bowl, emulsion side up. Pour boiling water over the top of the plate to completely cover it. The first time you do this, it is a bit nerve racking as you would imagine the glass shattering but its thinness dissipates heat and should be fine. Leave several minutes for the emulsion to absorb water, soften and expand. Using a plastic scraper (such as for clearing ice in a freezer), scrape the emulsion off. It will come off in great layers but will invariably leave some behind. It can be laid on a wet

absorbent sponge or kitchen towel and scraped again, or use a non-scratch scourer with the glass still hot with water. If the glass dries at all the emulsion quickly sticks again. The plate cools quickly but don’t be in a hurry to rinse under a tap; wait a few more minutes before rinsing. Rub dry and check all the emulsion has gone and if needs be, repeat the process. Once cleaned up, they can be dried and glued with silicone sealant to make observation cells. Leave in a warm place and dry for several days. When definitely dry to the touch, add water and leave for ten minutes or so to check for leaks. If you find any, dry thoroughly and cover the hole with fresh sealant. Dry gently with kitchen towel. Before using the observation cell, always clean the front glass panel with a window cleaner and paper towels. Clean and dry them after use. Keep observation cells stored carefully so they do not accumulate dust inside; in polythene bags is fine. After a few years the sealant may shrink and need replacement. Certainly after three to four years they begin to leak. Cut the sealant with a sharp knife and clean off old silicon, then rebuild the tank with fresh sealant. POND AND SEA WATER Aquatic creatures will need to be photographed in suitable water to meet their needs. If an animal will not settle in a dish or tank it may come down to the simple fact that the water is causing distress. Maybe the pH is wrong or there is a lack of oxygen. This can certainly be the case if you take water straight from the tap at home. I keep old plastic green bottles that originally held mineral water, filled up with water from different ponds and rivers that I use. There is also one with seawater. Before they are filled with the water it is fine filtered to remove as much organic matter as possible. That includes planktonic organisms. Filtered water is no guarantee that you have cleaned it but it will help the quality of the water to last longer. In addition, when magnified there is a surprising amount of ‘snow-like’ detritus floating about and the more you can remove, the better.

PREPARING SPECIMENS FOR BACK OR TRANSMITTED LIGHT This is a rather ambitious title as it suggests preparing material for the compound microscope, a potentially huge topic. Treatment can be quite specialized and we will deal with basic principles. References provide links to more information. Perhaps the most straightforward extreme close-up is the one where the specimen is held with reverse forceps and photographed with incident light. As you magnify the subject, so the density of the specimen can become a problem for showing detail. In Chapter 5, various methods were discussed that might help but ultimately if you want detail and the subject is too dense, it will be necessary to use a technique to thin the

specimen down to make it more translucent. Then it can be backlit in some way.

Fig. 6.17 Mouth parts of a rove beetle. The beetle was macerated in potassium hydroxide and eventually prepared as a permanent slide. Here, just the mouth parts have been photographed with a form of Rheinberg illumination. (×30 magnification. Canon 7D mk2 with bellows and ×10 Vickers objective lens, sub-stage lighting with 40-LED ring light. Shutter one sixth of a second, ISO 1600; composite of twelve-image stack.) Fig. 6.17 shows the mouthparts of a predatory beetle. At the magnification of ×30, even with the best incident light very little would be visible. By making the specimen as transparent as possible and using backlighting (in this case a form of Rheinberg) we see the structures revealed. Small material often benefits from being placed in water on a microscope slide under a coverslip, but a slight improvement will be made if instead glycerol is used to mount the specimen. This can impart some translucency as it has a different refractive index to the specimen. Glycerol-jelly is a step on from just glycerol on its own, and is typically used if you want a more permanent arrangement. Maceration is the principle way of making specimens soft and translucent by

removing the cell material and softening external skeletons. This can be used on just about any subject such as wood, insects and crustaceans. As maceration requires the removal of cell tissue, the process obviously has safety implications and needs to be done carefully and without splashing liquids. Have plenty of water available for washing anything that comes into contact with the chemicals or your hands. Potassium hydroxide is the chemical to use but sodium hydroxide is just as good and easier to obtain in the form of washing soda (often used to clear drains). A possible alternative for macerating softer tissue is a concentrated solution of biological washing liquid. The latter is not so good with tough external skeletons, such as beetles. Potassium hydroxide is purchased as a solid and it arrives in small pellet form. Play around with different concentrations but one pellet added to 30ml of water may suffice as a starter. Both potassium and sodium hydroxide are strong alkalis and must not get on skin, clothes or paintwork. If they do, wash immediately with plenty of cold water. Metal or plastic will not be affected. Like many of the chemicals mentioned here, treat with the utmost caution and ensure they are stored away from children and pets. Use of protective glasses and the wearing of nitrile disposable gloves is recommended. Make up a small amount of the solution in a container with a sealed lid. The solution cannot really be reused due to contamination and you may find yourself replacing the fluid after a day if it has gone dark brown. This is why only a small amount needs to be made up as required; it also goes off over time. Take the dead subject, such as a small insect. Note that the maceration process distorts wings if they are prominent on the specimen. Make a few holes if possible with a needle in the wall of the external skeleton (where it is not important for a later photo) to enable the liquid to penetrate more quickly and then leave to soak in a sealed container. The amount of time you need to leave the specimen varies, but could be several days. This could be accelerated by adding more potassium hydroxide – but be careful because if maceration happens too quickly, the entire structure can dissolve away. A thin specimen like a small fly will go quickly in less than a day but a tough ground beetle could be a week. Check fairly regularly to see how it is progressing.

Fig. 6.18 A freshly hatched moth caterpillar. (×16 magnification, permanent preparation with potassium hydroxide. HDMI inspection microscope, Rheinberg, composite threeimage stack.) During this time, the liquid becomes discoloured and needs an occasional gentle disturbance. When you think it is done tip the liquid into a dish, remove the specimen and wash in plenty of water. Putting the specimen in a series of tubes or small jars of water works well. This only takes a few minutes, then place on a slide. Try to arrange it as best as possible with a needle as the entire body will be flexible and soft. Cover it in water and place another slide on top. Press down firmly, holding for a minute or two. On relaxing, see if the specimen bounces back. If it does you may need to continue maceration. If the specimen is well and truly flat, the top slide could be removed and the water replaced with clean water and a coverslip. If the specimen will not lie flat when you release, one option is to replace the water, put a clean slide on top and fix it in place by wrapping tape around the ends of both slides to hold them together. By positioning the slide under a microscope or macroscope set-up, the specimen can now be photographed with transmitted backlight. This works very well with all insects and arachnids, making the skeleton soft and pliable. The effect is so dramatic that you may wish to make a permanent

preparation. You could mount it under a coverslip in glycerol or glycerol-jelly and seal the edges with nail varnish. This could remain viable for some years. The rove beetle in Fig. 6.17 was treated as follows: 1. After maceration, the taped slides squashing the beetle were placed in a jar of 4 per cent formalin to fix and preserve the specimen. It was left for one week. 2. The slides were separated and the now permanently flat specimen removed and placed in glacial acetic acid in a sealed tube for twenty-four hours. This removes water and dehydrates it. While you may associate acetic acid with vinegar, the glacial form needs to be treated with real caution. 3. The specimen is carefully removed from the acid and placed on absorbent paper to remove as much of the acid as possible before placing in clove oil. This clears the specimen of the acid and improves the refractive index further. Leave for twenty-four hours. 4. Remove surplus oil with absorbent paper and place on a clean slide. Add a drop of mountant medium; in this case Euparal was used but Canada Balsam is a good alternative. Place a coverslip over the medium. 5. Label and leave to set for around a week and this will last for many years.

Fig. 6.20 Head of tsetse fly from West Africa. With insufficient dehydration over time, water has clouded the specimen, surrounding it like frost. (×18 magnification, permanent preparation with potassium hydroxide. HDMI inspection microscope, Rheinberg, composite three-image stack.) All chemicals should be treated with care. For more methods and ideas see Peacock’s Elementary Microtechnique. For more details specific to arthropod (such as insects) methods, see Chick’s Insect Microscopy. We will come back to maceration in the final chapter. Of course, the reverse is true as well, that some material is too transparent to see any detail. Lighting such as dark field can generally deal with this. The traditional method of seeing detail in transparent subjects is to use some form of stain. This can be a general stain, like Eosin, that will dye the specimen red or one that picks up specific things like iodine, which turns starch grains a blue-black colour. Methylene Blue is another general stain that can make transparent objects a pale blue. Staining is a major subject in its own right and many agents can be administered to live material. HERE’S ONE I MADE EARLIER As a young teenager I loved microscopy but I was limited by my 8in tall microscope, which I had been given for Christmas. No doubt this is where much of my interest in extreme close-ups started. As a keen entomologist, my main concern was how to preserve and mount material so that I could see it under the microscope. Much later came the desire to photograph it. At the time

the specimens were made into permanent preparations and Fig. 6.17 shows a recent photograph of beetle mouthparts from a specimen mounted in 1967, the centre slide in Fig. 6.19. The permanency is not so important now as the digital image may suffice, although it does not take too much effort to continue the process and preserve the material for years to come.

Fig. 6.19 A sample of homemade prepared microscope slides.

Chapter 7

Focus Stacking?

any of the photographs that have appeared in the book so far have in their captions the words ‘composite’ and ‘image stack’. This refers to the way the photograph has been taken to attain the best possible depth of field by using more than one image of the subject and combining this stack to create one image; a composite. To remind us what was stated at the end of Chapter 1: as magnification increases, various issues occur including vibration producing blur, the need for more light and most important, the decrease in depth of field. Focus stacking is the answer as it produces a composite with incredible depth of field and is why the subject deserves a chapter to itself.

M

Fig. 7.1 Head of a young Water Stick Insect Ranatralineata found in pond water and photographed alive in an observation cell of water.×4 magnification, Canon 7D mk2, 65mm MPE lens with twin macro flash, ƒ5.6 ISO 100 composite of 34 images.

SHARPNESS AND DEPTH OF FIELD (DOF) The holy grail of the extreme close-up photographer is sharpness. This is not the

same as depth of field, which is more about focus and visible information. Sharpness is clarity and contrast to provide detail. Photographers know that when using diaphragm apertures or stops in a typical camera lens, the smaller the diameter of the hole (called ‘stopping down’ as it stops the light), the greater the DoF. Examples could be ƒ22 or ƒ32. By contrast a wide aperture such as ƒ2 or ƒ2.8 lets a maximum amount of light through but the DoF is very shallow. This information would lead us to believe that if we are having problems with DoF when getting close to subjects, a small aperture is essential. Unfortunately, while the DoF may be high the level of clarity can plummet due to distortion of the light passing through the aperture. This is all to do with the physics of light travelling in straight lines. Passing through a tiny hole like ƒ22 can cause the divergence of the light at an angle to produce a distorted image on the sensor. To improve this we look for a ‘sweet spot’, a particular aperture that has the minimum distortion. Usually this is a few stops down from wide open. Modern digital cameras can be confusing for beginners, as changing stops happens in partial increments, usually of thirds; here we are dealing only in whole stops. For example, if a macro lens has a maximum ƒ2.8, the next stop is ƒ4 and then ƒ5.6, ƒ8, ƒ11 and ƒ16. The sweet spot occurs around ƒ5.6 to ƒ8. The latter is more typical of full frame sensors and wider for APS-C. In Micro Four Thirds cameras, ƒ4 is often best. Ultimately, it is better to test this for yourself. Using only full stops, set up the camera on a tripod and in good, even light, photograph either a lens test card or some sharply defined printed text. Take a photograph on every aperture and then compare them, at least two side-by-side and enlarged on the computer. In this way you understand what apertures are suitable to use and which to avoid.

Fig. 7.2 Three photographs of a lichen growing on a fir cone, magnification ×4; each is a different aperture, from left to right ƒ2.8, ƒ8 and ƒ32. Note that the depth of field is very limited in ƒ2.8 and significantly greater in ƒ32. Note also that ƒ8 is the sharpest image. All taken under the same conditions with a 100mm macro with 120mm of extension tubes.

Fig. 7.2A The same as Fig. 7.2 but heavily cropped to show just the centre of the image. F32 displays huge distortion with no sharpness despite the greater depth of field. F8 is the sharpest image. Additionally, bear in mind that apertures may be irrelevant anyway. Many of the techniques needed to achieve high magnifications use lenses with no diaphragm present. Some manufacturers in the past have developed specialized micro lenses for extreme close-up, incorporating diaphragms with a small range of stops. Generally, these apertures have limited use. Test them but ideally use them wide open. A conclusion here is to aim for sharpness; depth of field is very limited whatever, and extending the DoF with focus stacking is the only option.

THE BASICS Focus stacking is often avoided by photographers because it can be perceived as a complicated procedure. The process can take time to set everything in place and get started, simply so that you understand the principles. To begin with try not to magnify the image too much, between ×1 and ×3 should be fine. Much higher and actually seeing the image can be difficult. What you will be doing is focusing on something at the front of the subject; take a photo and then move the camera forward a short distance before taking a second one. A slight overlap should be made between these two images. The camera is moved forward again before taking the third photo and so on until the end of the subject is reached. You then have a stack sequence, which needs to be combined on the computer to form the single composite. The ideal scenario involves checking this initial stack as something may need to be tweaked and the sequence run again.

Fig. 7.3 Composite photograph of a 32-image stack of bladderwort, Utricularia. It is an insectivorous plant, which lives in acidic lakes and ponds. If a water flea touches one of the hairs on the bladder it causes an in-rush of water, bringing with it the prey, which is caught and digested. A tiny, delicate structure that needed to be photographed in water. (×6 magnification, Canon 7D mk2 with 65mm MPE taken at ƒ5.6 with flash from behind and a fill-in at the front. Stacked with Zerene Stacker and edited to remove detritus from the water.) It is often the case that a stack has to be repeated several times. This is because with the front of the subject in focus, the back is invisible as the depth of field is so bad. Also light, even from something as consistent as flash, is inexplicably variable through a stack causing oddities that may require a re-run. To set up a basic stack you will need a focusing rail; in this instance, we will use a

manual one. Fix the camera to the rail and attach the rail to a firm support such as a tripod. For your first attempts, choose both a simple subject and simple lighting. Try to keep the stack on the horizontal as vertical stacking can have a few extra issues. With the camera near the middle of the rail, move the tripod close to the subject until it is nearly in focus. By using the rail, move the camera until the front of the subject is in focus. If in doubt exactly where this is, bring the camera back slightly until the picture goes out of focus. Using a remote shutter release or the self-timer on the camera, take the first photograph after a few seconds have elapsed from the last time anything was touched. Then move the camera forward to bring another area into focus. Again give a second or two before firing the shutter. Move forward and take another and so on, until the subject goes out of focus.

Fig. 7.4 and 7.5 Two photographs from near the beginning and end of the 32-image stack (Fig. 7.3) showing the very limited depth of field found in each one and why focus stacking is necessary. The imported images can then be checked for sharpness and the possibility of blur due to vibration. In the early days of your stacking, it is good to check the possible overlap between the photos in the sequence. Make a note of the interval distance that you use or if that is difficult with the manual rail, record the amount of turn of the knob on the side of the rail, for example a quarter or half a turn, and try to be as consistent

through the entire stack. Looking at one of the images at 100 per cent on the screen, quickly move to the next in the sequence while checking the same focus area on the photograph. Flicking back and forth, you can see if the overlap is sufficient. If not, make another run at the stack and check these. Once you have a known ‘interval’ between shots for particular set-ups, keep a careful record so that this degree of checking is unnecessary in the future. Produce a card or chart for future reference (see the table at the end of Chapter 3). Finally the stack needs to be combined into one composite, using software on the computer.

Fig. 7.6 Zerene Stacker screen in progress.

Fig. 7.7 Helicon Focus screen in progress.

STACKING SOFTWARE Photo editors like Adobe Photoshop and Corel PaintShop Pro, which use layers that can be combined, will work fine with simple stacks. However, if you find yourself running even a small but regular number of focus stack sequences, a dedicated programme is far better. The two principle players and without doubt the best overall are Helicon Focus and Zerene Stacker. Free software can be found but there is no comparison and little to recommend. Differences are minimal between Helicon and Zerene in terms of quality. They both produce superb results and are as good as each other. Helicon has the edge in speed (almost three times as fast) and is the most flexible in possessing a good retouching tool after the composite has formed. However, sometimes the extra speed of Helicon can create a few slip-ups in complex stacks. Also the composite can have more noise present.

Fig. 7.8 Zerene Stacker: part of the composite with no editing. Note the lines of dots and artifacts where detritus and other things within the frame have moved during the photographing process. This was cleaned up in editing and missing from the final result in Fig. 7.3.

Fig. 7.9 Helicon Focus: the same area of the composite as Fig. 7.8. Note the over-brightening and slight loss of detail where the highlights are blown. Figs 7.8 to 7.11 show a series of comparisons between the two. The bladderwort example is a delicate plant structure where individual cells are clearly visible, as the bladder wall is mainly one cell thick. Set up in a small observation cell, the lighting was with a flash from behind so the cells would be more clearly defined, with a slight fill-in flash at the front. The results from the stacking show the Zerene composite slightly better as the Helicon version has some highlight areas that are blown in a few small spots. Straight out of the stacker, the Helicon composite initially looks good but in this scenario, the Zerene has the overall better detail. Although both would require

some work in an editor, the blown areas would be difficult to recover. While Helicon took just twenty-four seconds, Zerene was slower at 1.52 minutes to create the composite.

Fig. 7.10 Zerene Stacker of a moth’s eye. Part of a composite with no editing. Notice in this case the loss of detail in dark areas.

Fig. 7.11 Helicon Focus: same area as Fig. 7.10 and in this case the detail is more obvious in the dark areas. By contrast, the moth eye in Fig. 7.10 and 7.11 shows the opposite. The detail present in the Helicon dark areas is better. Both again need some basic work in an editor but the Zerene composite needed more enhancement in the shadows, which created noise. Ultimately, the Helicon software had the edge with the result shown on the front cover. These are all fairly small points as both are very good and with some editing, the composites from both will be fine.

They both have more than one method to achieve the composite and nearly always, the best will be the pyramid or PMax. They need purchasing but (depending on which versions you subscribe to) give lifetime updates. Trial both of them and try different stacks to see if you have a preferred option. Despite having quite different interfaces they work in a similar way and maybe you will see that there is merit in having both! The stack of images can be added to the open Helicon or Zerene programme by either browsing and finding the files, or by just dropping them into the space reserved for the stack. For Helicon click the Render button or for Zerene click the Stack menu, then the top option Align and Stack PMap (or hold down the control key and hit P). The Zerene way is to choose the method each time as part of the start option. For Helicon you need to have selected the method A, B or C before clicking Render. The default is A, but by clicking C that will become the new default. As the programmes render the stack, the composite slowly forms in the window on screen. Upon completion, save the image. Both programmes display the output and you can stack any number of composites before saving, but probably best to check and save as you go. Both can be saved in either TIFF or JPG format although Helicon does have a further option for DNG.

TETHERING AND MONITORING IMAGES Until you become experienced at focus stacking, it is important to monitor the process with good viewing software. But even when you are highly proficient at the technique, an almost constant analysis of the images is vital. When you set up a stack sequence, before running it take a test shot to check lighting, sharpness and composition. With living material the subject can easily move during the stack, just a flick of an antennae or mouthpart. If this is discovered much later there will be difficulties trying to correct it on the computer. By collecting the stack of images on the computer as it occurs and producing the composite, a second run could be made when necessary. For an important stack it is worth running a repeat anyway in case of unforeseen issues like uneven flash lighting or vibration. The ideal arrangement is to use a computer connected to the camera during the stacking process.

Fig. 7.12 Focus stacking in the field. The camera is on a three-way geared head and connected by USB to the Windows tablet. The auto rail is a StackShot connected by the blue lead to the control box. The 12V battery and red/black lead power the rail via the box. The third cable from the box is the shutter-release lead connected to the camera. Fig. 7.12 shows focus stacking of life inhabiting the spaces between barnacles on a rocky seashore, a situation where there is no option but to take the equipment into the field. The camera, lens and lighting are on a Cognisys StackShot auto rail fixed to a tripod. This is connected to the control box, which in turn is connected to the battery at the back for power. The control box is also connected to the camera to trigger the shutter release. While live-view on the back of the camera is possible, the preferred option is to connect the camera by USB lead to the Windows tablet where Canon Utilities software can be used to set the controls of the camera in the photo. For other camera makes, the necessary software for the camera could be used or the alternative third-party software, Helicon Remote. The important issue is that live-view is now transferred to the tablet screen so that careful focusing is possible. In bright sunshine a coat is needed over the head and tablet so the screen can be seen. This was discussed earlier in the book under cameras when other options were mentioned. The larger live-view makes focus stacking easier, although it is largely about getting the stacks on to the memory card rather than producing a composite in

the field, which is completed later indoors. In the studio, tethering live view to a bigger screen is much more straightforward and the set-up would be similar. Every time a stack sequence is started you should check the lighting and exposure with a test shot. Again, looking on the back of the camera will help but viewing the image larger has to be better. The stack images should be quality checked, particularly in the field, as arriving home with many hundreds of images that all need deleting can be quite depressing. Check also that the interval between images in the stack show some overlap. Verifying images needs a simple but effective programme viewer. If you use Photoshop and Bridge or any other large, complex software you may struggle, especially in the field. After all, this is only about viewing and checking focus, not about editing images. There are plenty of free image viewers that can be downloaded, and many are small, requiring minimal RAM and storage. By having the image viewer running with the folder for downloaded photos open, the images can be checked as they arrive as the stack progresses.

RAW AND JPG IMAGES There is a general rule that using the RAW file in photography enables the most flexible and best quality final image after processing. The RAW data is taken from the sensor and no in-camera processing occurs, as this is reserved for the photographer to carry out later on the computer using software like Adobe Lightroom. In theory this is correct, especially if the processor knows what they are doing. Some cameras are better than others at in-camera processing and can produce excellent JPGs – to a large extent, this depends on the conditions in which the photograph was taken. If the lighting is good, such as a flash, and a low ISO has been set, avoiding noise as much as possible, then RAW may not be necessary. Focus stacking is considerably easier with JPGs rather than RAW as it is down to speed. Zerene does not work with RAW files and while Helicon does, this defeats the whole idea of using RAW. If RAW files are added to Helicon Focus, the programme has a RAW converter that automatically converts the image into a usable one. That can be useful if you have forgotten that the camera was set on RAW but as you have no control over the conversion, you might as well have used JPGs. The conversion process also takes time and so the stacking time will be around three times that of processing JPGs. The advantage of using RAW is when the lighting is difficult, such as a contrast subject with strong shadow or brightness or maybe when the ISO has had to be increased to beyond 800 when using LEDs. In these circumstances the RAW files need batch processing prior to the stacking. For example, when using Lightroom the images are imported so they are all present in a row. Edit and enhance the first one. When finished, select the whole stack so that they are highlighted and click Sync at the

bottom of the right-hand panel. This will bring all the files to the same state. Check there is not a rogue shot amongst them where the flash may have failed, or one that needed a different tweak. Of course, if that had occurred with JPG, the stacker would just have to deal with it but you have the opportunity here to do something with it. When all are satisfactory, export them as JPGs to a folder where the stacker can combine them. Another option (particularly with images that are just ‘noisy’ from high ISO settings) is to use DXO Pro, a piece of software that has arguably the best noise removal processing called Prime. Other adjustments can be set such as micro-contrast and exposure and then click to export all as JPGs. This is one of the least demanding RAW procedures as setting up requires just a few minutes and then hundreds of files can be processed in an automated batch. The drawback is that it is not quick but if you just want the computer to sort it out and you are not in a hurry, it works well. There is nothing wrong with working with RAW, but just expect that it is not going to be quick and only you can decide if it is worthwhile. Otherwise, JPG compositing is very quick to yield results. That is important if you are stacking a number of subjects and you need to know if the process has worked before you pack everything away. One of the big problems with magnification and stacking is that you can rarely see what you are actually doing. FASTSTONE IMAGE VIEWER This is my preferred viewer and although a Windows-only programme, I have several friends using Apple computers that have a Windows partition specifically to use FastStone. A free download of around 6 megabytes, it expands into an extremely useful viewer of all relevant file formats including RAW. Running in the background, it requires very little memory and stack sequences can be flicked through full-screen quickly to check exposure, focus and interval overlap. Up to four images can be positioned side-by-side and zoomed in and out of for comparison.

Fig. 7.13 Screen shot of FastStone Image Viewer.

DIFFICULT STACK SEQUENCES Fig. 7.14 is a composite of 117 photographs magnified around ×20. Setting this up and bringing the first part of the mosquito into focus for the first image of the stack concentrated on the tip of the proboscis. When looking at that tip, no single part of the rest of the head or body was visible. By moving the camera forward, the view changes as we run along the proboscis and the eye begins to appear and then the body. By this time, we are near the end of what will be the stack and probably realize that the subject is lying in the wrong plane and so needs adjusting. Invariably the specimen will need a slight rotation or perhaps moving side to side or up and down. This is why a moving stage is so important. When the specimen has been moved very slightly you then need to run back along the body, back to the tip of the proboscis only to find that that too is now in the wrong place.

Fig. 7.14 Mosquito composite stack of head and upper body. (Ú22 using a Canon 7D mk2 with bellows and Ú10 Vickers lens and twin macro flash seated on a StackShot auto rail.)

Fig. 7.15 Mosquito image number 5 in the stack sequence (Fig. 7.14).

Fig. 7.16 Mosquito image number 110 in the stack sequence (Fig. 7.14). Clearly, a flat object such as scales on the wing of a butterfly will be considerably easier, but these long stacks are worth the perseverance. Fine adjustment of the specimen may need to be done any number of times with you moving the camera back and forth to check the position of the subject for the start and finish of the stack sequence. Once the first initial stack of images is collected, it is important to check the composite to see if the end result is angled correctly. If not, try another adjustment and another stack sequence.

Fig. 7.17 Helicon focus screen while processing a 117-image stack of a mosquito head with the screenshot taken near the end. The window on the right is the developing composite while on the left is the photo being used at that moment by the software.

Fig. 7.18 The StackShot control box during shooting of the mosquito stack. We now see why an automated rail like the StackShot or WeMacro is so useful. With one click or hold of a button, the camera can be run back and forth smoothly to check the stack before running it. After all, if there are going to be hundreds of images involved, the process will be time consuming. There are two main methods that each of these rails employ to organize a stack and we will now run two stack sequences to show how this is done. Auto Distance is a method that sets the interval distance and Distance Mode is where the overall length of the stack is set.

Scenario 1: Using the Cognisys StackShot Auto Distance The mosquito was held in reverse-action forceps and it was most convenient to have this on the stage with the camera horizontal. Either of the auto rails could be used but the camera was fitted to the StackShot and bellows attached with a ×10 objective lens. The twin macro flashes were held either side of the specimen with simple diffusion, as reflection would not be a major problem. The StackShot moved the camera to a position around the middle of the rail and knowing the working distance for this lens to be around half a centimetre, the mosquito could be brought roughly to

that point. It was held by the wings and the position checked with a hand lens to try to establish a good viewpoint. It can be easier to see where the lens is located in relation to the specimen when held horizontally. By looking from behind the mosquito, the lab jack can raise or lower accordingly while the forceps can be rotated around until the position looks approximately right. The laptop was connected to the camera by USB 3 and the Canon Utility programme run, which enables the controls to be adjusted as well as calling up liveview. With such a limited working distance you can easily move into the specimen. Watching the screen carefully, the Back button can be operated first on the control box to pull away from the specimen. If you pass through the focus point, you know you are about the right focus. Then move forward and every so often, stop and check the distance from the specimen, until gradually it comes into focus. The stack was checked up and down to ensure the viewpoint and position were correct. Then a test shot was taken to check the exposure and position of the lights. Using the Canon Utility software, the flash exposure may be adjusted until that is correct. The control box was then used to set up the stack by selecting the mode. Auto Dist means the interval will be set between the shots and the first and last image set on the control. The Select button moves the left arrow on the screen (see Fig. 7.18) to select the row. With Mode selected, to move through the options press the Up and Down buttons. Various modes are available but select the Auto Dist mode. Then Select takes you down to the interval. There are some suggestions in the table at the end of Chapter 3 and there are various websites that provide ideas, but ultimately you will need to verify these for yourself, making sure there is sufficient overlap. For the mosquito stack with the ×10 objective, 0.02mm was chosen. Then Select takes you to the third row. This is where you choose the start and finish points. Using the Forward and Back buttons, the tip of the proboscis was found and then moved back slightly so it just went out of focus. The Up button was pressed (that sets the start point) and the Forward button moves the camera to the end of the stack. When you find this position, go just beyond where you want it in focus and press Up again (that sets the end point). The third time you press Up the stack will start. Before you do that the live-view window needs to be shut down on the computer or the shutter will not fire. Also you need to have configured the time gap from the StackShot moving forward and then firing the shutter, called the Settling Time. Two seconds should be enough time for vibration to stop and for the flash to have recharged. The capacitors in some flashguns can struggle with long stacks and if you find the camera shutter does not release, extend the time to four or even six seconds. This is done with the ConFig button but you only occasionally need to adjust these parameters. The control box will inform you how many shots it will take as well as progress. If you have the folder open on the screen within the image viewer, you can watch as the images arrive on the computer. You will be able to check the early shots, taking care not to shake anything. By starting just out of focus you ensure that the first possible

one in focus is included, as it is surprising what you might miss by mistake. Any auto rail will have ‘backlash’ or slack in the system. It becomes a problem when working with intervals of just microns, when on returning to the start point a rail overcompensates and misses the first shots. This can be allowed for simply by setting the start point way out of focus. Any early ones out of focus can be deleted and likewise those that come near the end can be removed, but stacker software largely ignores them anyway. When the stack has finished, the images can then be dragged and dropped from the viewer straight into the space in the stacking software. In the first place it is worth stacking all the images but when checking the composite, you may find that leaving certain ones out can improve it. Save the composite.

Scenario 2: Using the WeMacro in Distance Mode The WeMacro auto rail can also be used by setting the start and position but this scenario will use the method of Distance Mode whereby a set distance is programmed along with an interval length. Rather than give the start and finish position, just the beginning is found. As in the scenario with the Stackshot, the camera is connected to the computer so that live-view can monitor the process as well as adjustments made to the camera. The WeMacro auto rail, like the StackShot, has a control box but lacks any buttons or interface. Instead there is a USB connection through to the computer where the liveview is being monitored. Once all the hardware has been set up, the WeMacro control software is fired up. First click the Connect button in the window to make a connection to the rail. Using the Backward and Forward buttons, the subject can be brought into focus and checked. Take a test exposure and check to see if it is correct and whether the lights need moving to provide the optimum shadow and light areas. Also check for the need for diffusion.

Fig. 7.19 WeMacro computer control screen. A shows the one for selecting Start and End of stack. By clicking the By Distance box, screen B appears. This one is where the distance and interval are included. To programme the stack, find the start point by focusing on the foreground and then pulling it back slightly until the view is just out of focus. Remember that the WeMacro rail has two speeds: mm (fast) and micron (slow). Initial setting up will be easier in mm (fast) mode so select this from the down arrow and then save. The speed will be much quicker to find the start position, then move it back to micron

(slow) and save. Under Run mode, click the By Distance check point and if it is not checked already, click the ‘beep’ which notifies you when it has finished. The return check is to move the camera back to the original start position at the end so that it could be run again. As we will see it can be better not to do this in case you need to continue the sequence. Step length is the interval that needs to be added. A guide to this length is written in the table at the end of Chapter 3. For example, the MPE lens on maximum magnification with the lens at ƒ5.6 works well with an interval of 0.07mm or 70 microns.

Fig. 7.20 Vertical focus stacking with a WeMacro rail and stand. Note the silver control box, which is connected to a computer for the control screen. Ultimately you will need to try several options and look at the sequence full screen to check for an overlap in focus. Then a guess at the approximate distance the stack is to take is written into the Total Distance box. With practice, you will develop your own way of doing this but one suggestion is to aim for further than you think, such as 4mm or 4,000 microns as the stack can be halted at any time. As you add these two figures into the boxes on the screen, the Total Steps box changes as it calculates the

number that it will do. Now click Run and the sequence occurs automatically. If the camera is connected to the computer and you have set your preference for the images to automatically download, keep the download folder open so the photos are monitored and occasionally assessed to check that they are in focus. Once they start to appear out of focus, click the Stop button to end the stack sequence. Alternatively if the sequence finishes on its own and yet the last photo is in focus with more to go, then you need to continue the run. This is why it is best not to have checked the return after run otherwise the entire stack will need to be repeated. Instead to continue the sequence, the Run button just needs clicking again and the sequence will carry on for the prescribed distance. By assessing the images, halt (click Stop) the process once the images are out of focus at the end of the stack.

Fig. 7.21 Precision movement of the subject using a three-axis linear stage platform. WHICH IS BEST; AUTO DISTANCE OR BY DISTANCE? It is really down to you and your preference. I will use the Auto Distance 99 per cent of the time as I think it is more intuitive; after all, I determine where the stack starts and finishes. The problem comes with a long sequence at a high magnification. As we saw with the stack of a mosquito head when starting out, only the proboscis was visible and the end point tricky to establish. If you have set up an Auto Distance with a start and end, it is tricky to then decide when it finishes that you want it to continue. For those where the end is unclear then a By Distance option has an advantage as you can overstate the distance and get ready to stop the sequence as out of focus images appear.

MOVING THE SUBJECT While moving the camera is the usual default method of focus stacking, if the specimen is small and not going to run off anywhere then there are some advantages in keeping the camera still and moving the subject. Success does depend on subjects that will not be disturbed by moving, such as dead creatures or aquatic organisms in cavity slides. As long as they can be fixed in some way, this can be the preferred method if you have heavy photographic equipment, especially when operating the camera vertically. By the time you have a DSLR with a lens like an MPE and macro flash, you may be reaching the limit of the auto rail. Using the flash off-camera will help and bellows with objective lenses will be fine. The Zeiss Tessovar is a superb piece of kit if you can source one, but the weight with a DSLR can be too much on an auto rail. In these cases, fix the camera and lens in position and use a method to move the specimen. A focusing rail, whether manual or auto, can be adapted quite easily to hold a platform or crocodile clip to mount the specimen. The rail moves the specimen just like the scenarios above. If you are using lab jacks for moving the stage, you might want to try them to move the specimen vertically towards the lens but they do not necessarily travel smoothly or in an exact straight line, as there can be lateral deviation. If you want real precision use a sliding table, around 6 × 6cm, called a 3axis linear stage platform. They can be moved in any plane and with an accuracy of just 0.01mm. While not cheap, one might be useful if you want to manually focus stack to this precision. A fairly straightforward DIY platform can be made that will rise vertically by using a long screw thread and several guides fitted vertically to a base, then threaded into the movable platform. One poor result with moving the specimen is possible change in lighting during the stack. Helicon Focus can sometimes come up with a red triangle on the file name of a composite and invariably, an inconsistency occurred with lighting, giving a percentage. Over a small distance this should not occur and if a message appears over a long stack, the composite can still be fine.

STACKING ON A MICROSCOPE Focus stacking was initially developed to help the problem of shallow depth of field in

microscopes. The way that most people do this is by focusing the fine adjustment to just above the specimen and then take photographs after a slight adjustment downwards on the fine focus. Watching down the microscope or on live-view, the travel may be as little as a tenth of a turn on the fine focus adjustment.

Fig. 7.22 Homemade microscope stackers attached to the fine adjustment knob. Left: made by Ray Sloss to fit a Wild M20 and based on the stacker, right, constructed by Mike Crutchley for a Zeiss. For regular stacking you might think of setting up an automatic device to assist. WeMacro sell a simple set-up for doing just this, fitting on to the fine focus knob. Fig. 7.22 shows two homemade devices, both based on similar principles. They are fitted to the fine focus knob at the side of the microscope. By pressing down on the lever at the bottom and side, the stacker rotates the knob. The degree of travel can be set by the brass wheel at the back. As the lever comes down, the knob rotates and a microswitch sends a message to the camera to take a photograph via a cable release. Simple, but very effective in making stacks.

USING VIDEO FOR STACKING With a flick of a switch, modern digital cameras can go from taking still images to amazing quality video. Ignored by many as a gimmick or simply for very occasional

use, video clips can be used to create difficult stack sequences. A typical frame rate (number of images taken per second) is twenty-eight or thirty. This means that a 10second clip can have around 300 images. Individual frames can be extracted and used, but the quality until recently was quite poor. A few years ago this improved somewhat with HD quality forming 3 megapixel images but it was not until 4K video arrived in cameras that frames of 8 megapixels made extracting images worthwhile for stacks. The only real problem with video is that it tends to waver slightly from side to side. One of the great improvements in stacking software in recent versions has been the ability to align the images to create the composite. Auto rails also maintain excellent alignment. The consequence of this is that now the use of video to produce a stacked image has never been so good. To do this, move the camera on the rail while watching live-view to the beginning of the stack, set the video running and then move the camera until the end of the stack. Stop the video. StackShot tends to run a little too quickly while WeMacro has the advantage in micron mode to run more slowly. Helicon Focus deals well with video clips. Just drop the clip into the space and the software starts to extract the frames and set these as individual files in the top right. At the end you see the number of frames present and many will be useless. At the beginning there will be a big delay in the camera starting the video and moving off, and again at the end. In fact, it can be best to set a time; for example, count two seconds after switching on so that the equipment settles down from having been touched before starting the movement of the rail. If that is two seconds, there will be around sixty frames at the start that will be useless. Click on the sixtieth frame in the sequence and check to see if it is worth using. Do this on a number of files until you find the ideal one to start the stack. The ones not being used need selecting and removing or the tick removed. As well as the beginning, there will be a number of frames to remove from the end of the stack and switching off the video. Then click Render. Using video works well as long as the movement of the camera is smooth and slow. Why use video? Stacking with stills is the preferred method as the process is more controllable, but using video is a good standby when speed is of the essence; for example, in the field with a live specimen that will only stay still for a few seconds. Track the organism with the camera and the moment it stops moving, a short clip is taken as the camera is moved slowly forward. It does require practice to get it right. Try breathing (a subdued and gentle blow) on the moving creature as that sometimes stops them for a few moments. Crucially this technique does not require critical focusing as the video can be started before it is in focus and then while it is running, the camera can be moved. Just be selective over the frames that are stacked. The results are often better with lower magnifications with a macro lens rather than bellows and objective lenses. If ambient lighting is good, video clips can work well with difficult stacks and this is only going to get better; 4K is now available at 60fps

and the Lumix G9 has 6K video, allowing 18mp images to be extracted. VIDEO FOCUS STACKING ON MICROSCOPES If there is one area that using video clips to generate composite images is particularly useful, it is when using a microscope. It is also very straightforward. The image below is of a polyp on a marine hydroid colony magnified ×100. The subject is three-dimensional and transparent. Colour has been produced by using a special lighting technique on the microscope called differential interference contrast (DIC). More important is the difficulty in creating a good focus-stacked composite as a high number of images will have to be taken, adjusting the focus each time. This will take several minutes and the chances are the subject will move, albeit slowly. Using the fine focus, the top stack position was found and put just out of focus. As the video recording on the Sony A6500 was started, so the fine focus knob was slowly rotated to move the focus point down the specimen until it was out of focus and the recording stopped. Lasting six to ten seconds, each clip was then imported into Helicon Focus, creating an average of 250 frames to stack. In many cases, although only 8 megapixel images, they are better than a manual stack. Taken on an Olympus BH-2 microscope.

STITCHING COMPOSITES There are times when a specimen will not fit to the photo format or the possible magnification; the specimen just cannot fit the picture. Either the specimen is too big to fit the field of view of one magnification and too small for another. This is often the case on microscopes where there is a limited range of objectives and do not forget that the sensor area or photograph will be an enlarged part of the actual field of view.

The problem with Fig. 7.23 is that the array of diatoms was too large for the use of a ×10 objective and ×4 was way too small. Secondly, a specific phase contrast objective to produce more contrast was required and was only available as a ×10. A further complication was that the diatoms on the slide are quite three-dimensional and so needed focus stacking.

Fig. 7.23 Diatom composite made from four separate composite stacks that were then stitched together.

Just over a quarter of the diatom arrangement could be photographed at a time with the ×10 objective. Starting with the top left quarter, a ten-image stack was made. Then carefully overlapping, the top right quarter was photographed, again with a ten-image stack. The subsequent bottom right and bottom left stacks were made. The composite for each quarter was made so that four images could now be stitched together to create one photograph. This was done using Microsoft’s free ICE (Image Composite Editor) programme, when the four composites were dropped into the programme space and the software combined them in both planes. This is then exported into a photo editor where the final image was given an improved background and trimmed to make a photograph with a magnification equivalent to around ×80.

Fig. 7.24 Head of a greater water boatman, Notonecta (×3). This was made in a similar way to the diatom composite by making four separate composites and then stitching them to form one. Not only is there more detail, but it was the only way to do this with bellows and ×4 objective; even with minimum extension, there was too much magnification.

Fig. 7.25 Marine flatworm; four images stitched horizontally to form one image.) Objects that do not move, like the preserved diatoms, are clear-cut and with care this can be done with material that rarely stops moving. Expect the need for patience and a low success rate although with experience it is possible. For example, the flatworm in Fig. 7.25 is a tricky customer. The primitive creature belongs to a group found in freshwater and on the seashore. They are not true worms at all and the tiny body seems to glide across the substrate, never stopping still as the flat base is covered in minute hairs called cilia, which beat to create the movement. They are abundant – the example here was found amongst some seaweed and is just 3mm long. There is a head end to the body, despite there being no brain, so you do know which direction they will be travelling. The flatworm was transferred to a cavity slide and immersed in seawater, with a coverslip over the top. Regardless of having added magnesium sulphate, it was moving fairly fast. Under a bellows with a ×4 objective lens, the flatworm still moved too fast to grab the entire body. Twenty or so photographs were taken, moving the slide on an X–Y mechanical stage to try and track the animal. Flash was used to freeze action but none of the photos had the whole animal, only sections of head and mid-body and tail. Ultimately there were four images chosen which had at least one section of the body sharp. Two of these needed cropping or straightening so that on adding to ICE, the pieces would join up. The result was not perfect and a small amount of cloning was required, followed by a cleaning-up of the debris scattered around the animal. A winter species of mosquito that visits our houses at this time was photographed

to show the amazing detail of the head. The specimen was dead and upon looking at the composite, although the mouthparts looked spectacular, they were less so out of context with the rest of the body. The answer was to produce four linear composites with plenty of overlap between them and then stitch these composites into a panorama. The outcome was an image with appreciably more impact, showing what looks like an imbalance of structures on a small head and body.

Fig. 7.26 Male winter mosquito, Culiseta annulata. A: a composite image of the head to show the elaborate antennae and mouthparts. B: four composites are stitched in a panorama to show a more dramatic image of the head appendages against the context of the body.

FOCUS STACKING PROBLEMS The mosquito composite in Fig. 7.14 did not stack exactly as the image shows. In fact there were some major problems and artifacts (visible defects) that needed dealing with. Depending on what you are stacking, as many as one in ten composites can have one or more artifacts. A common problem is that as you move the camera,

areas change size and if the camera moves at a slight angle along the axis of the stack, odd lines appear around the edge of the picture. These are easily removed by cropping later but will result in a lower resolution image. If there are some unwanted effects of the stack you could try running the software stack again but change the method, such as B instead of C (Helicon), DMap instead of PMap (Zerene). If you have both software packages try the other one, although current versions have become very similar. For long stacks, say over a hundred images, they can be broken down into smaller groups. For example, if the stack is a hundred put them through the stacker in four batches of twenty-five. Then stack the resulting four composites. Sometimes that works.

Fig. 7.27 Composite problems of Fig. 7.14. These are the original composites created by A: the pyramid method and B: the mapping method.) The mosquito stack resulted in several strange gaps. Most strange was the proboscis which, instead of joining the head in the correct place, stopped at the eye and appeared to come from the back of the head. The stack of images was split into six batches, each with plenty of overlap. The result was better but the base of the antennae and the exact point of contact for the proboscis was still not good. To

improve moments like this, both of the software packages have retouching tools. Essentially this is the same as cloning as found in photo editors, but the difference is that the files are listed in order on the screen and the composite next to it. This means that the clone areas match with no setting up required. What you do have to do is to find the files with the sections you want to clone on to the composite. Click through the files and each time they will appear on the left of the composite, so you can check if the missing piece is there. In the case of the mosquito it was the base of the proboscis first and so, with such a limited depth of field, it needed a small piece of material cloned from around fifteen files to retouch the image. If you have created four or five composites by batch processing, you may be able to reduce this to cloning just one of the composite pieces rather than finding fifteen separate files. Retouching is relatively easy and the results are clearly improved.

Fig. 7.28 Close-up of the problems in Fig. 7.27. Sometimes the only answer to a strange and significant artifact is to repeat the photographing of the stack and reduce the interval distance between the images. The most common problem to occur during the taking of a stack is for something to move. This will result in multiples appearing in the composite. Plants can have quite surprising amounts of movement. The scarlet pimpernel photograph was repeated several times due to the central stamens moving during the stack; tiny seeds inside seed heads absorb water and sometimes move during a stack; worst of all would be the close-up of the reproductive structures under fern fronds. The intriguing structures

are like lollipops that have spores inside. To disperse them, they absorb water and expand under warmth (such as lighting) and then flick the spores into the breeze. Live animals will flick a mouthpart or antenna. These movements occur often so subtly that until the stack is done, they go unnoticed. With care, most can be rectified by retouching in the stacker but this is another reason why repeating a stack sequence several times is good, so that you have the options later. Even what appears to be a perfect stack can have a few issues that need dealing with, such as detritus. This is obvious when stacking specimens in water. During the photographing of the stack, tiny pieces of debris sink down through the liquid. Picked out by flash, the small dot appears in each shot but not in the same place as it is sinking. On the composite, this one piece of detritus can be replicated a dozen or more times and in turn can be multiplied by the number of pieces occurring. As they can run in lines of dots they will be distracting and ideally removed. While this can be done in the stacker retouching mode, it is probably easier to do this at the editing stage along with other aspects needing attention.

Fig. 7.29 The centre of a scarlet pimpernel flower (×5). The stack had to be repeated and retouched due to movement of both the pollen and stamens under the lights. Dust on the sensor can be a problem for all genres of photography where it creates darkened patches in light areas on photographs like the sky, but with extreme close-up photography, the patches become more dominant, coming sharply into focus. With a stack sequence, due to the movement of the camera, this in-focus dot now forms a curved line of small black dots that will require removing at an editing stage.

Composites created with the pyramid method tend to increase noise and even purple fringing. Colours may be muted and in need of a boost in vibrancy and clarity. When composites look like they need work in an editor, they should be saved in TIFF format to avoid the affect of compression. Several lenses, including objective lenses when on bellows and especially extension tubes, can result in darkened corners due to vignetting. All of these can be adjusted in programmes like Lightroom, DXO or Capture One as well as normal enhancements such as Levels, Curves and Unsharp Mask.

FINAL POINT WITH FOCUS BRACKETING We saw in Chapter 2 how Olympus have incorporated focusing bracketing in some of their cameras, even the excellent TG compact series. Setting up focus bracketing results in a series of images from the foreground to the background where the camera has adjusted the focus of the lens rather than by moving the camera. The images are then used in Helicon or Zerene to produce the composite. The Helicon software company in 2016 developed a small extension tube, which fitted on to Canon or Nikon bodies and to this you fixed a lens, such as a macro. The tube contains some clever electronics that you programme on a phone app and then transfer the settings across wirelessly, such as interval and aperture. By using the continuous shoot mode on the camera, you hold the shutter release down so the tube adjusts the focus between shots to create the stack sequence. The Helicon FB tube works well after initial teething problems with the app and data transfer to enable focus bracketing to be used automatically in more cameras than just the Olympus. While this is a good method of focus stacking, it relies on a focusing lens and has limited use in extreme close-ups. SCORPION SCENARIO Illuminated with just a single torch of ‘black light’ the aim was to show the brown creature under 365 nanometres of ultraviolet. The scorpion was a pinned specimen held about 20cm above a black sheet of card. The camera fitted with macro lens and extension tubes on a manual rail attached to a tripod was set up with LED spots in a darkened room. The focus was set at the nearest point. Then the lights were switched off, protective glasses put on and the torch switched on. By playing the light on to the scorpion’s head, the first shot was taken a sixth of a second at ƒ8 with ISO 400. The rail was moved an eighth of a turn and allowed to settle for two seconds before releasing the shutter again. This was repeated until the scorpion was out of focus. Due to the poor lighting conditions the files used in the sequence were RAW to maximize the quality. The stack sequences were repeated several times with bracketing of the exposure. On the computer, the stacks were renamed in FastStone and sorted into their sequences. Taken stack at a time, the first image in the stack was enhanced in Lightroom (mainly exposure, contrast, clarity and noise) before exporting as JPGs to be stacked in Helicon Focus. The composite was then put back into Lightroom for minor tweaks including import into Topaz DeNoise and then Detail (all as TIFF files). The images were finished off with a bit of cloning in FastStone to remove the

bright patches that had burnt out, the pin and shadow at the top and two parts of limbs on the right that seemed distracting.

Fig. 7.30 Scorpion composite photographed under 365nm of light. A is the original composite and B is after work in different software.

Chapter 8

Inspiration for Extreme Close-up Photos

is in fashion when the gorse is in flower’ is an old country expression ‘Kissing indicating that whatever the day or season, you will always find gorse in flower. So many photographers pack up their macro gear as the country enters the throes of winter, thinking that photo pickings become slim. If anything there are more weird and wonderful creatures about; the mosquito we saw at the end of the last chapter appears late in the year. Discovering material is a little more of a challenge but much of the life will be original, as far fewer people will have photographed it. There is always interesting, natural things to find displaying super texture, colour and pattern.

Fig. 8.1 Autumn is a time for wild fruits and seeds. On the outside they may look tattered and dull, so look inside and at the seeds. Interior of the marsh marigold (Caltha palustris) seed pod with membranes displaying a beautiful texture, like old wood, with seeds in pockets. (×6 magnification. Soft light created with cone diffusers on twin macro flash, Canon 7D mk 2 with 65mm MPE lens.) The aim of this chapter is twofold: suggestions through the year of what might be about and could make good extreme close-ups and, secondly, by choosing some

specific examples we can look at ideal methods to photograph them. The list here is minute compared to the practically infinite variety of possibilities. It just requires time and a hand lens to explore what is out there, as it will not be long before you will be photographing something no one else has. Finding out what it is might be more of a challenge!

AN EXTREME CLOSE-UP YEAR Seasonal change brings the chance to find new and varied flowers and non-flowering plants. Like gorse, red campion and members of the geranium family can be found on every day of the year in the British Isles. Stinging nettles always make good images and they can be found in one form or another at any time; it is just their shape and thickness that change. Yet another reason to keep photographing the sting cells around the stem and leaves. While out on a walk, whether along hedgerow or through woodland, go slowly looking around you for subjects. A hand lens is fairly essential so that the material can be examined closely to see if it has potential. Carry small containers and plastic bags. Intricate detail of the flowers is good but by taking some home, like gorse and broom flowers, they can be dissected in a dish and the tiny insects living there extracted. Thunderbugs are such a group of insects; they are just a matter of a few millimetres long and feed by sucking out the contents of cells. Ponds and streams are good for most of the year. In the summer much of the insect life is present as adults in the air above the water. Eggs are laid and hatching occurs in late summer and early autumn. This results in quite young stages and so at these times a pond can seem a bit empty. Development of the young stages increases until the onset of winter, when it stops. Rather than hibernate insect larvae enter a diapause period, which means growth is suspended but they remain active so they can still be found. Early spring sees the development continue until they mature. This is a simplification of the larger animals’ life history and plenty of the smaller creatures like gnats and mosquitoes can be present and emerging throughout the year. The main insect emergence period commences in April through to September, although many small insects continue throughout the year.

Fig. 8.2 Hover fly larva eating an aphid. Aphids themselves are interesting to photograph but finding a mass of aphids will have plenty of different predators in attendance. (Canon 7D mk2 with Raynox 500 fitted to a 70–200mm lens set at ƒ16, flash; six images stacked and retouched as the gut moved.)

Fig. 8.3 Hazel, Corylus avellana, female flower, early spring. The male catkins are obvious while the female flower requires a search along the twig as it is so small. ×8 magnification, Canon 7D mk2, bellows with ×4 objective lens, twin macro flash, composite of forty-image stack.) March to June is the period of maximum density of freshwater and marine plankton. A smaller peak occurs in the autumn but something will always be found swimming, particularly in the sea. Plankton from both fresh water and coast can be collected with a fine mesh net of around 100 microns (60 microns is best but more expensive). This will collect various tiny crustacea like the water fleas and copepods. Along the coast you can add much more in the way of crab and barnacle larvae, not to mention the larval stages of all the shore creatures like periwinkles, limpets and fish. Towing a plankton net behind a boat at a slow rowing pace is ideal but a small net on a long handle can be moved along the edge of a jetty or pond. Try to avoid large sections of weed.

Fig. 8.4 Marine plankton, the larva of a sea urchin. (×100 magnification, composite of four images, dark field lighting.) Even when the frost is thick and snow covers the land, head for the coast because the seashore never fails to deliver on any day of the year. A quick glance at a rocky shore seems to show a constant source of large seaweeds and the usual array of periwinkles, limpets and other life. Certainly these creatures are always there and easy to find but there is also a seasonality on a finer level, with a huge abundance of small, delicate seaweeds developing in the spring and summer. Breeding of seashore

animals varies through the year – do not be surprised in October if tide pools become white as the limpets discharge their sperm and eggs into the water. Search under seaweeds and amongst the weed that fringes the pools. Collect a small handful of these fringe seaweeds, place in a pot with seawater and take them to a home studio to tease out in seawater for tiny animals. Finding life on sandy beaches is more difficult, as the animals live between sand grains and are buried. Open sandy coasts will be poor in material as the waves shift the sand and prevent animals settling. Enclosed, sheltered sand and mud shores are always very rich in microscopic life. Scoop up soft samples near the surface to examine under a stereomicroscope with the sediment in a small dish of seawater. Transfer animals to cavity slides and dishes to photograph. Sediment shores include salt marshes where small pools form at low tide in which tiny creatures can be found. When visiting any place with soft sand, collect a small sample to magnify at a later date. Most times it is silica particles, while occasionally it can surprise with exquisite mollusc shells and even those of a group called foraminiferans.

Fig. 8.5 Head of a shield bug. (×8 magnification composite image.) Here are a few specific ideas with approximate timings, as their presence will depend on latitude: Spring (March–May): the mud and sand of salt marshes, collect samples in the pools and mud banks; compost heaps for worms, snail eggs and insects like springtails and so on; flowers in woodland and hedgerow; insect pond life; insects emerging; plankton; grass and tree flowers. Summer (June–August): insects of heathland; feathers; pond life; moths through to

autumn; flying insects generally peak at this time; cow dung for insects and fungi; delicate seaweeds, including the life amongst them, growing on larger seaweeds mid to lower shore (wash large seaweed in a bucket of seawater and animals emerge, which can be collected and taken to the studio). Autumn (September–November): many spring flowers and creatures can appear again like violets, cow parsley, geranium family, foxgloves and plankton; seeds and fruit; ferns, especially their underside with reproductive structures; fungi, especially in fields and woodland; marine mud; reproducing lichens and mosses. Winter (December–February): early flowers like snowdrops and late fungi in woodland; under the bark of dead trees and inside well-rotted logs – many insects like beetles, solitary wasps and other invertebrates hibernate here; ivy flowers from late autumn, angle-shades moth; seashores are always a good prospect at this time both for strandline and rock pools for fringe seaweed with organisms; some pond life. For further ideas and identification aids, there is a series of small books called the Naturalist Handbooks listed at the end of the reference section.

Fig. 8.6 Close view of a fallen, rotting pine tree with much of the bark removed. Holes, burrows and sawdust can be seen where the tree has been eaten by invertebrates. There are numerous fungal threads visible. This is an ideal source of specimen material in winter as many insects from the surrounding area come to join the existing community to hibernate inside the log.

COLLECTING MATERIAL I have an old fishing bag with a variety of plastic tubes, jars and containers of different sizes. In the base of the bag is a white 2-litre ice cream box along with sealable plastic bags. When collecting samples with a net from ponds, rivers and rock pools, the net is emptied into the ice cream box and sorted with pipettes and plastic white spoons. Also in the bag is a collapsible butterfly net. Additional nets include two different pond nets, with either a short or long handle. The short handle can be removed and fitted to a different net called a sweep net. This has a triangular opening at the front and rather than netting, it is made of white callico. This net is swept back and forth through vegetation and creatures drop off into the bag, which can be searched with a tube to collect specimens. The bag has a pocket that neatly takes a UV torch for searching at night for scorpions and other arachnids, although it is surprising what glows or shows a different colour in the dark. I rarely need to carry the bag far as I always have too much in the way of camera gear, so the fishing bag and nets are in the car and used when needed. In addition, a moth trap is great to leave out at night for a collection of material for the following day. Insects other than moths are attracted including flies, caddis flies and ichneumon wasps. Small yellow dishes with soapy water in them can be placed outdoors which will attract tiny insects that can be extracted dead from the water to photograph. Plastic yoghurt pots sunk into the ground with the lip just beneath the soil surface make good pitfall traps to collect ground invertebrates.

SPECIFIC PROJECTS Previous chapters have given a rather one-sided approach by tackling a specific aspect of the technique in taking extreme close-up photographs. This section will suggest eight different subjects that you might not have considered before and look at particular issues that affect the photographic process. A wide variety of areas have been chosen to follow through background information, problems and equipment. 1. Texture using fruit and seed Often, what makes a good image is one that others cannot see with the naked eye or where there is a difference in scale and perspective. The photograph that started this chapter is one of my favourites as, despite a huge magnification, the subject looks like an old, gnarled piece of wood. The image has a familiarity about it but the actual material is a thin membrane inside the fruit of a marsh marigold just a month or so old. Examining one of the dark fruits on the end of a stem with a magnifying lens, it was cut in half and the membrane could be seen with the seeds. It was set up on reverse forceps and photographed with flash angled low to the surface. Texture is an important element in photographs and one we should look for in the micro-world. With still life, texture is straightforward to set up. From July through to November there is a huge variety of seeds worth looking at, either with or without the fruit which contained them. Most have an unusual surface to them and a shot we could set up here is with red campion seeds. A very abundant species in hedgerows

and grassy waysides, the fruit is a dark brown flask shape that opens at the top to release large numbers of tiny black seeds. Viewed with the naked eye, there is nothing to make you think it is worth photographing. Collect a number of the fruits complete with a small amount of stem and check that they are not empty. Place one in a holder such as reverse forceps with the opening slightly angled to one side. Add more seeds from another fruit so that the seeds are bursting out of the top. Set the camera horizontally with a suitable lens. Here, I used the 65mm MPE lens, but anything magnifying in the region of ×3–5 will work. Focus on the lip of the fruit and expose for the seeds. They are virtually black but have a super texture creating a monkey-like face. To achieve the texture, light – in my case a flash – is held out to one side of the subject and used to skim a bright light across it so the roughness of the seeds forms a shadow. A second, weaker source of illumination is on the other side. Alternatively, use a reflector. The image is created by stacking a series of images across the opening.

Fig. 8.7 Seeds: red campion. (×5 magnification.)

Fig. 8.8 Seeds: tiny seed picked up on socks in the outback of Australia. (×6 magnification.) Seeds are very light and move easily. During the stack of Fig. 8.7 one of the seeds moved slightly and had to be touched up with cloning. Small piles of different seeds can produce lovely patterns. Use a hand lens to inspect the surface of different seeds to look for unusual textures. A small area near the base of the wing on a sycamore seed is like burnt wood; the inside of the fruit of beech mast can be quite beautiful; birch seeds are wind-blown and collect in large drifts like dust, but go in close and the fine, delicate wings with backlighting are very attractive. 2. Colour and texture on insects In the same way as seeds, a magnified section of insect parts can reveal fabulous texture and colourful patterns. Almost any moth or butterfly wing reveals stunning patterns in the scales. A close-up of just an eye can disclose unusual patterns, most notably in horseflies. One of my favourite parts to photograph is the underside of beetles or where the wing cases join at the top of the body. Often they can be beautifully metallic while others are deeply pitted, which creates interference patterns of light. To display these textures, treat them like the seeds before and skim light across the surface. Wing scales can need high magnification and require the use of bellows fitted with a suitable objective.

Fig. 8.9 Insect colour: left, scales of a green arches moth and right, close-up of the eye of a clegg (small horsefly), both ×20 composite images photographed with bellows and objective lens. 3. Water mites and other moving, aquatic animals Fresh water, such as ponds and lakes, has an excellent variety of microscopic, planktonic creatures that can be abundant in the spring and summer. Many species of water flea, or daphnia, can be found and Cyclops, a copepod. After netting them and sorting through a sample under a stereo or HDMI microscope, they can be extracted with a pipette and placed in a drop of water on a cavity slide. Carefully add a coverslip by placing one edge on the slide and lowering the other side gently with a needle so that large air bubbles do not get caught. If they are moving too much, they can be slowed with a concentrated drop of Epsom salts solution placed on one side of the coverslip with the drop in contact with the water in the cavity. A piece of tissue or kitchen towel is put on the other side of the coverslip to soak up the water. As it is absorbed, so the narcotizing salts are drawn into the cavity. Do not be tempted to do this too quickly or it may kill the animal rather than just slow it down. Dead animals have a tendency to take on an unnatural look. Water mites are a little bigger at around 2mm and trickier to deal with, as they tend to be fairly immune to Epsom salts. These tiny arachnid species exist in a wide range of shapes, patterns, colours and size. One thing for sure is their inability to keep still, which makes them one of the most difficult of all aquatic groups to photograph. They have a bulbous body and eight feathery legs that never stop moving. Instead of the drop of water in a cavity slide, fill it with a drop of wallpaper paste. It probably does not help that much but I usually add a little Epsom salts to the paste as I make it up. With the water mite in the pipette, try to squeeze the specimen out with as little water as possible, not in the paste but somewhere else on the slide. Then using a needle, coax the animal out of the water drop and carefully into the paste-filled cavity. With the needle, mix it slowly into the paste, trying to avoid introducing air bubbles. The

mite may remain near the top of the paste but that is fine, as you need to lower a coverslip on top and push gently down. The water mite continues trying to swim but very much in slow motion. The main point is that the animal is not going anywhere and so the slide can be placed under a vertical photographic set-up.

Fig. 8.10 Fast-moving pond life: a water mite photographed in wallpaper paste. This will have been prepared beforehand and in the example shown here, the camera, bellows and objective lens were mounted on a vertical auto rail. I always have an old permanent prepared slide handy so that I can get the focusing point ready for when I put the cavity slide underneath. It saves a good deal of searching around and requires only a small amount of refocusing. It is unlikely that the water mite will completely stop moving and that will prevent a full stacking scenario. Instead manually change the focus and take photos of the body and some of the legs. A flash

may be necessary to freeze action so the legs are not blurred, as they are likely to move at least slightly. A twin macro flash was used with one on top and one below the specimen to provide backlighting for the hairy legs. As long as there are at least two photos on the computer, the sharp in-focus body can be easily cloned on to an image that has good detail in the legs. One image alone of the legs in focus will have an out-of-focus body and this can be dealt with by cloning. In the example here, the paste did not mix well, creating an odd background in the original image and as there was also detritus introduced with the mite, the background was replaced on the computer.

Fig. 8.11 Pond life: saucer bug head from beneath, three composite images stitched. (×4 magnification.) 4. Small invertebrates during mid-winter By far the most popular extreme close-up images that photographers enjoy taking are of insect heads. Several reasons stand out: ease of capture, availability of material and the unrivalled diversity of insects. The ideal portrait of any invertebrate will be the head and maybe part of the body. There are a number of these across the course of this book. Unfortunately as autumn moves along into winter, most terrestrial life shuts

down. However, there is a great habitat available the year round which is especially good during the winter. In fact, the photos taken for this section were taken one day during early January after a thick frost had frozen every other part of the country. Decay generates heat and keeps the invertebrates active. This works with a wellprepared garden compost heap but the very best is nature’s largest compost heap, a habitat called strandline found on the coast. In the British Isles it is said no one is more than seventy miles from the sea, but the best strandlines are found along a coast where seaweed grows. During the autumn gales, huge piles of seaweed are wrenched from the rock and deposited at the top of the beach. These are continually added to by each subsequent tide throughout the year. Within weeks this immense mass of organic matter rots down to be recycled back into the sea. The organisms that create this unequalled decomposition vary from micro-organisms through to invertebrates that consume this unending arrival of organic material, to the predators which feast on beasts beneath in the food chain. What makes this particularly special is that these creatures are unique to the strandline and rarely leave it.

Fig. 8.12 Strandline: seaweed fly, Coelopa. (×4 magnification.)

Fig. 8.13 Strandline: head of a borborid fly that is just 2mm long. (×16 magnification.) A mixed sandy and small shingle beach not too far from a rocky shore where seaweed grows may be worth your first visit during the winter. Take a few large bin bags and a pair of thick gardening gloves. Be aware that glass, plastic and any amount of potentially sharp objects can be washed in with the seaweed. Also as the seaweed is eaten and decayed by the invertebrates, there is a release of the slime (called algin) found in seaweed. Larvae (maggots) of the flies love this bath of nutrients but it is messy, so gloves make life a little more pleasant. You need to get down to the base of the ‘compost’, move to the side any recent deposits and go for the richer material on the sand or shingle. Most obvious will be the crustaceans like sandhoppers that start jumping; this is the layer that needs scooping up in your hands

and putting into the bags. Once you arrive home, open the bags slowly and a little at a time. Be ready with tubes and containers to catch and sort the invertebrates as they appear. Many flies will emerge and so doing this in a garden shed or garage may help prevent escape as they fly to the windows. Beetles, including a few weird-looking rove beetles, should be abundant. Some of the smaller insects like the flies are fascinating up close. Examine some of the slightly unusual maggots as their breathing holes, located at the rear end of the body, are raised up to stop the slime entering. Some of the small specialist beetles like Omalium tend to collect in the bottom corners of the bin bag so do check these out. Remaining strandline material can be kept in the bag, tied off and left outside in the cold for a few days so that more specimens can be removed as required. When finished with, what remains can be added to any compost heap where it seems to boost the quality of what was already there!

Fig. 8.14 Strandline: head of a rove beetle from the strandline. (×8 magnification.) Once collected the specimens can be placed in the freezer to kill them prior to photography. Reverse forceps are good for holding the insects or setting them up on some of the decaying seaweed. Entire rove beetles look better if they are photographed alive. This is possible by keeping them in a fridge for an hour and then putting them in a small observation cell suitably attired with seaweeds. They need a minute or two for any condensation to disappear. I have also photographed maggots in this way. In all the cases here, the camera was set up horizontally either for the

observation cell or forceps in the case of the flies. Twin macro flash was also used with the light angled at 45 degrees. 5. Transparent subjects – photographing mollusc tongues or radula This may seem to be rather bizarre subject matter but actually they are quite beautiful and intricate structures. There are plenty of small transparent specimens that could have been selected like hydroid colonies growing on seaweed or phantom midge larvae found in ponds. Instead this is an example of the many amazing hidden natural structures. Nothing is easier than finding a snail or slug although some effort will be needed to extract the tongue. A characteristic feature of all gastropod molluscs is that to be able to feed, they have a long strap-like tongue (the proper term is radula) with suitable teeth, grooves or extensions on it to rasp plant or flesh. In most cases the tongue is quite transparent and there will be some difficulty in seeing the detailed structure, hence our choice of subject. But first we need to find the radula. Common pests in the garden, slugs and snails will work although they are more varied and different to those found on a rocky seashore. Even a brief visit to a rocky shore at low tide will reveal periwinkles, topshells, dogwhelks and limpets, each with very different tongues. Pop a few of each type (shore or garden species) in a jam jar and then keep them in the freezer until you need them. Thaw them out before use and then you need to break the snail shell (except in limpets). Garden snail shells break easily after freezing but the shore molluscs need more persuasion (a hammer generally does it). Freezing also minimizes the amount of slime although marine species do not cause a problem. With much of the shell gone, place the specimen in a dish covered with water so you can identify the key parts. Along the base of the body, including the head, is a muscular foot. Above this are the main body parts and above that is the digestive gland (which can be removed). The radula is quite long and located at the head end, just in from the mouth. The limpet is the easiest, just cut with a sharp craft knife into the muscular foot near the head and the tough radula normally emerges as a dark loop. Using a pair of forceps, pull it away and you may be surprised that unravelled, it is almost the length of the body. Other species are not so visible as the radula is more transparent. The best method is to cut away the head and front section of the foot and then place this in a tube with a dilute solution of potassium or sodium hydroxide to macerate the body. A strong solution of bio-washing powder works as well. The tongue is tougher than the surrounding tissue and as that slowly dissolves away, you should be left with the long, thin, strap-like structure. This is usually hours rather than days but the process does vary with species, for example garden snails require far less time than shore species. If you leave it too long the radula will dissolve as well so tease apart with needles as it dissolves, looking for a long ribbon structure. Do not let your skin come in to contact with the chemical at any time; use

forceps and keep plenty of water nearby in case of contamination. Some, like the topshells, are not so ribbon-like but more the shape of a long boat. With the exception of a limpet, a radula on its own lacks contrast and colour and requires careful lighting to display the delicate structure. Dark field illumination is a good start or use Rheinberg to introduce some colour. In the example here for a periwinkle radula I have used a stop system, like Rheinberg, but instead of the stop being blue and black it is blue with a yellow centre (Fig. 5.17). A red and blue one was more colourful with contrast, except it created a rather garish result.

Fig. 8.15 Radula: set-up for photographing the radula on a stop box in Fig. 8.16. A: the specimen on a slide; B: large convex lens from an enlarger to direct the light up to the specimen and C: the blue and yellow stop under the lens.

Fig. 8.16 Radula of the edible periwinkle. The periwinkle radula was cleaned in water and then added to some dilute glycerol on a slide under a coverslip. The glycerol prevents the slide from drying out as well as slightly improving contrast. The set-up is shown in Fig. 8.15 and a WeMacro auto rail held vertically over a special box stage. The slide is at the top of the stage, receiving light from below. Inside the box is a powerful LED and above it is a 20mm diameter hole with the blue and yellow filter held in place with a convex lens. This directs the light up to the specimen. Without it, the light is too scattered. Attached to the auto rail is a DSLR fitted with fully extended bellows and a ×10 Vickers objective lens. All of this is on a heavy table on a wooden floor and therefore subject to vibration. I decided to stack in manual. The WeMacro software was set at an interval of 15 microns. Using the back and forward buttons on the software, the beginning of the stack (the nearest area in focus) was found. Tethered to a laptop, this is easy to see in live-view. The Canon Utility software was running for the live-view to appear but neither WeMacro nor StackShot can fire if live-view is open and so the shutter was released by clicking the Canon Utility button on the screen. By doing this with liveview, the Canon camera will use an electronic first curtain shutter that reduces shutter vibration. Following this first photograph, the Single Step button is clicked on the WeMacro screen to advance the camera. When all vibration around me had stopped, I fired the shutter again on Canon Utility. And so on until the screen went out of focus. To reduce blur as much as possible a higher ISO was used than when using flash and so the photos were RAW files. The specimen is not moving and with no rush to complete the stack, RAW is fine. These were converted in Lightroom by enhancing

the first image, including removing the vignetted corners, and then the rest of the stack sequence was synchronized with it. After exporting these as JPGs they were stacked in Helicon Focus to create the composite image. The final photo was tweaked slightly in Lightroom and Topaz DeNoise.

Fig. 8.17 Radula of a common topshell using crossed polarized light; composite, stacked on a M20 using the Ray Sloss stacker, Fig. 7.22. (Photographed by Mark Papp.)

Fig. 8.18 Fungi: eyelash fungus. (×5 magnification, 65mm MPE, composite of a 42-image stack.) 6. Fungi Everyone is familiar with the classic toadstool but most fungi are macro and microscopic. Fig. 2.1 is a close-up of a pin mould growing on a tomato in the fridge. The kingdom of fungi exhibit extreme diversity; the list of species in Europe run to many thousands and new species are being found almost daily. They are constantly around us even though we associate autumn with fungi, as that is the time when the reproductive structures appear after a windfall of organic matter. The textured surface up close is interesting although the underside and a close-up of just one of the gills can be better. A small piece of the spore-developing gill in glycerol under ×40 can show up appealing structures.

Fig. 8.19 Fungi: black tar fungus on a sycamore leaf. (×25 magnification with bellows and ×10 objective, composite of a 58-image stack.) When out walking, always keep an eye out for small coloured spots on the ground where organic matter might build up or on bark, especially where decay is spreading. Carefully extract the fungus by removing part of the substrate or if possible complete a stack in place. The eyelash fungus is quite common on acid moors and is visible as just a tiny orange spot on the ground. Bird’s nest fungus forms on the side of grasses. Fungi can appear anywhere at any time. Marine fungi are very small and may occasionally turn up on old seaweed. While compost heaps will have fungi, one amazing place to explore, not just for fungi, are cow pats in a field. So astonishing are they that Peter Skidmore wrote an entire book called Insects of the British Cow-dung Community. Excellent for beetles and flies, there are some beautiful fungi too. The cannonball fungus produces spores which need to pass through a cow and grow in

the cow pat, but to pass back into another cow the fungus has to shoot the spores well clear of the cow pat to reach fresh grass, in order to be eaten. Their glass-like appearance needs careful diffusion of the light.

Fig. 8.20 Fungi: edge of a small toadstool. (×6 magnification.)

Fig. 8.21 Kelp at low tide on a rocky shore; a holdfast is ringed. The black spots that appear on the leaves of sycamore late in the year are from black tar fungus. Only very close examination shows the fungal threads growing across the surface. Fully extended bellows fitted with either ×10 or ×20 objective will show up the fungus and the consequence of the infection. In the photograph here, I have used a single flash from the side to create the texture of the leaf surface. To stop any movement and ensure the leaf is as close to 90 degrees to the lens as possible, small magnets held it tight to the metal bars on a ‘helping hands’ device. 7. Kelp holdfast If you are netting for plankton along a marina, always look along the edge of the decking that is permanently in the water, allowing kelp to flourish. Alternatively on a low spring tide the kelp is revealed in the lowest areas of the rocky shore. Using a knife to prize them off the substrate, first search the long fronds for anything useful that might be attached and then cut these off to leave just a small piece of stalk and the base, the part we are after, called the holdfast. This can be sealed in a plastic bag before returning to the studio. The dome-shaped holdfast is a mass of root-like branches that provide a home for many hundreds if not thousands of creatures to hide

within the spaces. Tiny worms (Fig. 5.1) crawl out as well as sea spiders and small crustaceans. Attached to the holdfast surface will be small anemones, star ascidians and hydroids.

Fig. 8.22 Kelp holdfast: tiny crustacean called Idotea.(×5 magnification.) Pull the holdfast apart in a white dish of seawater to flush out the creatures. Using a fine paintbrush and pipette, transfer the specimens to cavity slides or small dishes whilst larger material will need placing in small observation cells with a little seaweed. Fig. 8.22 is a young Idotea crustacean, commonly associated with seaweeds. Using an observation cell with an opening at the top of about 5mm, some seawater was added that had been filtered first. Filtering will remove most of the detritus that would otherwise cause a ‘snowstorm’ of particles in the water. The cell was on a lab jack and the camera and lens set up horizontally on a StackShot with twin macro flashes either side of the cell. The flash prevents blur if the animal moves. The live creature did move slowly along the seaweed and a series of shots were taken. The three chosen photos would not align properly due to the movement and so they were merged manually. The best example (the one with most in focus) was chosen as the main image and then the better sections on the other two images were selected and pasted on to the main one. This could be done by cloning.

Fig. 8.23 A: Kelp holdfast: small gammarid. (Original photograph.) B: final photograph of small gammarid image after changing the background. The smaller creatures in cavity slides with coverslips and seawater included a little Epsom salts to slow movement. The little gammarid crustacean in Fig. 8.23 is normally very active, but slowed sufficiently to allow a fifteen-image stack to be completed. The cavity slide was set up on a glass stage, beneath which was a black background. The vertically mounted bellows and camera had a ×4 objective lens, a flash was used from below and a fill-in on the top. Unfortunately the background was

very distracting, as the flash has emphasized cleaning marks on the glass. To improve this, the creature was cut out in Topaz Remask and replaced with a colour selected from the original background.

Fig. 8.24 Kelp holdfast: a common but rarely seen species of crustacean called Tanais. Note the two barnacle larvae found in the plankton. Photographed in a cavity slide with wallpaper paste made up with seawater. (×4 live composite of four images) The second example here was a crustacean that is common but rarely seen, called Tanais. Photographed in a similar way to the gammarid, this animal did not keep very still and so of the sixteen images taken, three separate composites were made up of three, five and six pictures respectively and the rest deleted. Then a final composite was made of the three composites. At each stage a small amount of retouching in the stacker software was required. The two small planktonic creatures accompanying the Tanais are two nauplii larvae that appeared in several of the shots and are included in the final result for size comparison. The diverse material found in a holdfast could keep you going for a very long time. 8. The surface of leaves We end with something that seems mundane and yet has an incredible variety of form and texture. Some, like the stinging nettle and bladderworts, we have already seen. The more extreme the environment, the greater the chance of a strange structure to the leaf. Sand dunes and saltmarsh are particularly interesting as the leaf surfaces have special glands to excrete salt and trap moisture. On moorland and heaths you will find the weirdly adapted leaves of insectivorous plants like the bright red tentacles

of the sundew species or the microscopic stalked globules on butterwort. Whenever you are out for a walk, take a hand lens and just look. Photographing them is straightforward and diffusion of the light is often necessary.

Fig. 8.25 Leaves: sand dune plant showing thick hair for trapping sand grains. (×4 composite of 36-image stack completed in the field, Canon 7D mk2 with 65mm MPE.)

Fig. 8.26 Leaves: the top surface of the insectivorous butterwort, Pinguicula, showing a digested gnat among the stalked globules of mucus and enzymes. (×6 composite of 55-image stack.)

References and Further Reading

Websites Brunelmicroscopes.co.uk Source of microscopes and equipment. Cognisys.com StackShot autofocusing rail. coinimaging.com Excellent website with lens tests and comparisons. Edmundoptics.com/uk/ Equipment and information. Extreme-macro.com Information and reviews. Heliconsoft.com Stacking software. Mccrone.com Mainly microscope-based tutorials and information. Microscopy-uk.org.uk/ Information, magazines and used equipment. Photomacrography.net Excellent forum for information and help. Quekett.org/ Main site for the Quekett Microscopical Club; information and resources. Rms.org.uk Royal Microscopical Society. Srb-photographic.co.uk Adapters and filters.

Tethertools.com Software for tethering cameras. Watdon.co.uk Watkins and Doncaster for field collecting equipment and moth traps. Wemacro.com Autofocusing rail, lenses and equipment. Wwb.co.uk Worldwide Butterflies for equipment, moth traps and live specimens. Zerene.com Stacking software. Books Blaker, Alfred. Handbook for Scientific Photography (W.H. Freeman and Co., 1977). Bracegirdle, Brian. Microscopical Mounts and Mounters (London: Quekett Microscopical Club, 1998). Bradbury, S. and Peacock, H. Peacock’s Elementary Microtechnique (Edward Arnold, 1973). Chick, Andrew. Insect Microscopy. (Crowood Press, 2017). Davies, Adrian. Digital Ultraviolet and Infrared Photography (Focal Press, 2017). Galtsoff, Lutz, Welch and Needham.Culture Methods for Invertebrate Animals (Dover Publications, 1957) – an old book but with a huge amount of information about growing, feeding and sourcing material. Grey, Peter. The Microtomist’s Formulary and Guide (Original 1954 – now printed new on demand). Lawson, Douglas. Photomicrography (Academic Press, 1972). Matsumto, Brian and Roullard, Carol.An Introduction to Digital Photomicrography (Crowood Press, 2017). Ideas and ID Books– Naturalists’ Handbooks, a series of small paperbacks. These thin paperbacks are full of ideas on how to find abundant small invertebrates and microscopic organisms in common places and have been produced by a series of publishers over the years from Cambridge, Richmond Press and now Pelagic Publishing. Excellent second hand copies are available through portals such as www.abebooks.co.uk. The following are a few titles that may stimulate ideas and help with identification: Animals on Seaweeds by Peter Hayward (1988).

Animals of Sandy Shores by Peter Hayward (1992). Insects on Cherry Trees by Simon Leather and Keith Bland (1999). Animals under Logs and Stones by Philip Wheater and Helen Read (1996). Insects on Nettles by B. Davis (1991). Insects and Thistles by Margaret Redfern (1995).Insects on Dock Plants by David Salt and John Whittaker (1998). Microscopic Life in Sphagnum by Majorie Hingley (1993). Animals of the Surface Film by Marjorie Guthrie (1989).

INDEX

Photographic Topics adapters 26–27, 43, 45, 47, 62–63 auto distance (in stacking) 118–121 autofluorescence 87 backgrounds 95 backlash, autorail 120 battery 29, 31, 49, 50, 69, 70, 78, 115 bellows 27, 36–37, 42, 44, 46, 48 black light 7, 87, 131 block cameras 29, 54–57 bridge cameras 29, 37 bright field lighting 59, 60–62 Canon MPE lens 41, 49, 122 cavity slides 99, 140, 154 chromatic aberration 24, 38 clamps 94 compact cameras 27, 38, 61–62 condenser, sub-stage 60–61 copy stand 92 coupling 8, 29, 37–38 CSCs/mirrorless cameras 25–26, 31, 35–37, 41–42, 49, 54 dark field lighting 11, 60, 78–80, 90, 148 depth of field 13, 15, 17, 46–47, 61, 109–110 detritus problem 98, 129–130 diffusion and diffusers 67, 70, 73, 75–77, 79, 95 distance mode (stacking) 120–121 DSLR 23, 31, 49, 54 epi-lighting 83–85 epsom salts 100, 140–141, 154

euthanasia of specimens 98, 146–147 extension tubes 35–37, 40–42, 44 Faststone Image Viewer 116, 131 flash diffusers 73, 76 direction 70, 138, 152, 154 macro 72, 146 ring 73 using 67–70, 79, 120 flat field 33–34, 43–44 fluorescent light 85 fluorochrome 88 focus bracketing 27–29, 130 focus stacking 25–26, 28, 30–31, 46, 69, 109–131 auto distance/by distance 118–121 difficult sequences 117 microscopes 122–123 problems 127–129 focusing rails 47–51 focusing 18–19, 26–27, 29, 33, 40, 47, 59, 141 auto/manual 18 fringing 34 glycerol 100, 103, 107, 148 Helicon Focus 112–118, 123, 127, 130, 131 Helping Hands 94–95, 152 ICE (Image Composite Editor) 125 Köhler illumination 60–61 lab jack 94 Laowa lenses 41, 46 LED lights 35, 78–79, 86–87, 94, 116, 131 lens extender 40 lenses, achromat 34, 44 apochromat 4, 44 bellows 42 convex/concave 33–34

enlarger 42–43 planachromat 34, 44, 64 live-view 19, 23, 25–26, 29–31, 47, 63, 118, 120–121, 149 maceration 103–105, 147 macro converter 38–40 Magic Arms 92, 94 magnesium sulphate 100 magnets 76, 95, 152 magnification 11–19, 33, 35, 38, 42–44, 51, 58, 60, 83, 103 chart 17 problems of 17 methylcellulose 100 microscope, clip-on phone 21 compound 53–54, 58–61, 83, 88, 106, industrial HDMI inspection 29, 55–57, 81, 96, 107, 140 stereo 53–55, 57, 96, 136, 140 USB 56 mirror lock-up 23 narcotizing specimens 99, 140 noise 13 numerical aperture 44 objectives, 34–36, 43, 63 finite 44–46 infinite 46–47 oblique lighting 80 observation cell 39, 69, 97, 98–103, 113, 146, 154 optical bench 92–93 parfocal 63 peak focusing 26 phase contrast light 64–65 photomicrography 9 photomicrography 9 photosites 13 polarization, cross 81–83, 148 Polymorph plastic 73 potassium hydroxide 104, 147

preparing specimens 103 Quekett Microscopical Club 63 RAW files 27, 115, 149 Raynox lenses 19, 38–40 reflection 68–70, 74, 77 refraction 33 resolution 21 reverse forceps 94–95, 138–139, 146 reversing lens 37 Rheinberg illumination 80, 83, 103, 148 Royal Microscopical Society (RMS) thread 45–46 sensor dust 19, 130 sensors 13, 16, 21, 23, 25, 27, 34 sharpness 109–110 shutter actuations 25 shutter, first curtain 24, 26, 149 smartphones 21–23, 27, 61 snoots 70 sodium hydroxide 104, 147 softbox 73 spherical aberration 33–34, 43–44 stacking software 112–114 StackShot autorail 49, 78, 92, 115, 117–121, 123, 149 stage for specimens 93–95 mechanical 95 three-axis linear 122 stage micrometer 13 stitching images 125–126 stop box 81, 88, 146 stops (lighting) 79–81, 148 tablets 30, 50, 115 tethering 29–30, 114–115, 149 texture 70, 138–140, 152 tripod 91–92 UltraViolet 7, 85–89, 131

vibration 19, 23, 25–26, 30, 33, 67, 69, 120, 149 video stacking 123–125 viewfinder, right-angle 48 vignetting 23, 34, 45, 61, 149 voltage regulator (Safe Sync) 72 wallpaper paste 65, 99–100, 141 WeMacro autorail 49–50, 117–121, 123, 148–149 stand 92, 94–95, 120 working distance 33, 35, 38, 40, 44–47, 63 Yasuhara Nanoha lens 42 yellow filter 88 Zeiss Tessovar 41, 122 Zerene stacker 112–118, 127, 130 Zhongyi lenses 42 Index of Species Photos alga Phycopeltis 84 anemone, beadlet 28 snakeslock 28 barnacles 28 barnacle larvae 154 bee 14 beetle, click 75 Donacia 15 great diving 98 rove 81, 104, 145 water (larva) 102 bladderwort 31, 111 Bosmina, water flea 65 Bowerbankia 102 butterfly, clouded yellow 71 green hairstreak 11 butterwort 155 diatoms 52, 124

Phaeocystis 90 dragonfly, gold-ringed 74 flatworm, marine 126 fly, borborid 144 doli 77 dung 45 empid 50 horse 96, 140 house 89 hover (larva) 134 predatory 40 seaweed 143 tsetse 107 fungi 35 marine 9 fungus, black tar 150 eyelash 149 pin mould 21 harvestman 19 hawkmoth, small elephant 108 hazel female flower 134 hydroid 87 Idotea (crustacean) 152 kelp 151–155 lichen 110 limpet 28 louse, head 37 marsh marigold seeds 132 mosquito 117, 127–128 moth, green arches 140 onion, wild 77 opossum shrimp 11

periwinkle, edible 147 phantom midge (larva) 62 plankton 64, 135, 140 Portuguese Man o’ War 99 prawn 97 pseudoscorpion 8 red campion 138 saucer bug 142 scorpion 6, 131 sea mat 102 sea urchin (larva) 135 shield bug 136 spider, crab 36 tarantula 86 stinging nettle 66 Tanais (crustacean) 154 tintinnid 64 topshell 148 wasp, emerging adult in nest 68 pupa in nest 76 ruby-tailed 16 water boatman, greater 126 lesser 69 water mite 141 worm, (marine) 32, 39 leaf (larva) 90 sand mason (larva) 99 spirobid 101