Prism Projection

I’ve written a bit about the DynaZoom and DynOptic before, I’m almost sure I have, but today I thought a bit about where the stand really shines; teaching.

DynaZoom & DynOptic in the Classroom

The stands of the Dyn* generation aren’t my favorite, the fixed inclination is just something I never understood, even in the BalPlan it’s irksome. There is, however, one thing to which the Dyn* line is particularly well suited and that’s instruction. Simpler U or bird-foot stands have a tendency to be roughly handled by students, and particularly in the case when used with a mirror in the light-path will almost certainly never been in alignment, not so with any of the Dyn*’s. The heavy base sits still and because the illumination is integral (unless used with the rather rare-mirror base) will remain aligned provided it is properly set up once.

They’re also great in that of the various optical heads, the photomicrographic tribute-nocular is enormously common. Although it was available with any number of camera bodies, 4×5, Type 80 Land Camera, Polaroid pack film, even now 35mm remains the most often seen. The comparatively (these days) outdated film cameras provide an excellent jumping off point for someone wishing to adapt a digital camera to the microscope. One could still stumble upon the somewhat rare B&L C-Mount video camera tube and teach a class with a single microscope if one had a mind to. But the right angle prism eye-piece is more common, and can be applied to other stands as well.

The Prism Eyepiece

B&L, perhaps more than any other microscope manufacturer had accessories. There was seemingly a device for every need to be had and at least here, outside of Rochester, New York, many of them still turn up on the yard-sale and thrift-store circuit. The B&L prism eyepiece is one many a microscopist would do well to pick up.


B&L right angle prism eyepiece

It’s a simple two-part thing, black enamel body bearing a right angle prism in an adjustable mount (the angle of movement is only 10 degrees or so), and a friction fit collar for the eyepiece tube. The collar pulls out from the body and slides easily over most standard 1/16th wall eyepiece tubes where a tiny knurled set screw secures it to the tube. With that in place an eyepiece is installed as normal and the body of the prism eyepiece slipped onto the collar over that. Anyone with more than the most limited experience with B&L illuminators will have noticed that most light sources they made provided a range of illumination that may be described on a scale of too-bright to I’m suddenly blind.

Obviously most illuminators they made we’re meant to be used with skylight or neutral density filters even on their lowest power for visual work. With the prism eyepiece it will be clear just why they provided such ample light.


There’s a lot to be said for the utility of gazing up from the eyepieces for a large and clear view projected upon a wall or screen. Even excluding pains in the neck, it can be Wonderfull for taking notes, or even simply for giving the eyes a bit of a rest, the projection set up for some distance can be a nice way to exercise the focus of ones eyes while not interrupting the use of the microscope.


Distance to wall, some 27 inches

Light Source Power Supply Alternatives

Get an amazing deal on a stand and have everything you need except the power supply? Don’t leave it on the shelf in the hope of one day converting it for an alternative bulb (LED conversion can be great, it can also be terrible, don’t rush into it) or wait for the day to come when an appropriate transformer turns up, just buy an autotransformer! A suitable autotransformer won’t exactly be cheap but can prove quite economical in the long run, we’ll get to that later, first lets look at what one is, and what’s normally provided. First a little bit about power, lights, and dimmers.

Most of the time, in a residential application that is, light bulbs provide a constant fixed level of brightness (marked on packages in lumens) but generally thought of by the consumer in terms of watts. A watt is a description of energy that is equal to the voltage multiplied by the amperage. The conventional 60w incandescent light bulb may be powered by 120v at half an amp of current, or 12v at 5a. Years ago if one wanted to get a lower level of illumination from an incandescent bulb one used a dimmer switch that contained a variable resistor that limited the voltage which traveled to the bulb the current remained the same. We can see then that the same bulb which provided 60w of illumination at 120v and half an amp would provide 30w at 60v and half an amp. With these type of dimmers the energy that is restricted by the dimmer (30w) is dissipated by the dimmer as heat, no energy is saved, between the dimmer (30w) and the bulb (30w) one is still using 120v and half an amp.

More modern dimmers operate by means of a simple circuit that rapidly turns power traveling to the bulb on and off. It happens so fast that it’s invisible to the eye and with incandescent bulbs the heat held by the filament makes the fluctuation even less noticeable. The advantage of such a dimmer is that there is some minor reduction in power consumption—the reduced wattage output by the bulb is not dissipated by the dimmer as heat. Unfortunately, a by product of such a dimmer is a reduction in the working life of the bulb which should avoided like the plague in the case of frequently difficult to find and expensive to replace microscope illuminator bulbs. Both of these dimmer types, both the old fashioned and modern have one significant flaw for the microscopist—in a word current—but more on that later.

An autotransformer is and electrical voltage transformer of a special sort, functionally a dimmer of the old-fashioned type described above, yet instead of functioning like a resistor it’s a transformer and functions due to induction. Unlike a standard transformer with a primary winding of a particular number of turns and at least one secondary winding of a differing number of turns, an auto transformer has only one winding. In a normal transformer the supply voltage is connected to the primary winding and is output at the secondary at a different voltage that can be calculated with a set of equations.

If the primary has more turns of wire than the secondary the input voltage produces a lower voltage on the secondary it’s called a step down transformer. If the situation is reversed and the primary has fewer turns than the secondary (or the secondary from the first example is used as the primary) it’s a step up transformer and outputs a higher voltage than was input. This is how the old, heavy, wall adapters are able to output 12v even though the socket on the wall provides 120v or 240v.

An autotransformer isn’t automated or automatic, rather it’s so designated for the fact that it is self-transforming. In place of two separate windings a single continuous winding is used for both the primary and secondary. The use of a single winding means an autotransformer rated for a particular input and output voltage will be much smaller than a standard transformer with a primary and a secondary. With most autotransformers the primary and secondary are not in a fixed permanent relationship, but are variable across a given number of steps.  One might just as easily be continuously variable across a range. Most are able to provide an output from significantly lower than the input voltage to a bit over, though others are constructed specifically to provide much higher output voltages than those input. For the purposes here we’ll want an autotransformer which takes a standard input voltage and can output a range from < 1 up to > the input voltage. The practical application of such a device is that an significant number of different bulbs can be run from a single transformer rather than needing a different unit for any particular microscope.

At most hardware stores a standard lamp dimmer can be had for as little as ten dollars, throw in a box in which to mount it and a hardwood base and the whole deal still only just approaches twenty bucks. A secondhand autotransformer might turn up for $50 but one is better off buying new where one can find a 120v autotransformer rated for 20a (of the cheapest sort mind you) for around a hundred, more than five times the cost of a dimmer switch. An autotransformer as described above even if rated for only 10a might weigh as much as twenty or thirty pounds. The reason an autotransformer is preferable has to do with amps and volts. A dimmer switch will only work at the rated voltage, but that’s still not the worst thing about them, after all many student microscopes from the 1960’s and even the 1980’s used a 15w 120v night-light style bulb, it’s a question of current.

The dimmers at the local hardware will at most be rated for 4a, maybe a few for high voltage halogen track lights go as high as 5 or 6a. That’s more than enough for a single incandescent drawing even 2 to 3a at the most. Now, something like the B&L Dynoptic that takes a GE-1634 only draws a single amp, a touch more if over-run to 25v, so a small resistive dimmer would do if installed after a step-down transformer, but it wouldn’t be very efficient. That same bulb could be easily be run by an autotransformer, place a tape mark or two on the control knob to a avoid accidentally feeding it a drastic over voltage and you’re in business.

Where the autotransformer really stands out however, is when it comes to running much older lamps from much different types of illuminators. The first incarnation of the B&L Research Illuminator dates to the early days of electricity and took a range of bulbs from 120vAC to battery based 24vDC home electrification systems that were in use in rural areas for decades before rural electrification ramped up the late 1930’s and post war 1940’s. The second and re-designed Research Illuminator (the model with the rectangular horseshoe base) took as standard a flat filament incandescent that was rated for 18a at 6v. The original power supply for the 100w bulb was about the size of a breadbox and looked and acted much like an early electric space-heater.

The all-metal units contained a large step-down transformer and a multi position switch that would remove one large resistor from series for each step the switch was moved to increase voltage fed to the lamp. It might seem strange that the unit simply didn’t employ a number of secondary windings and so provide a range of voltages with a single component. The use of the resistors made the unit smaller and cheaper to manufacture. Some workers would strip the switched resistor series and replace it with a rheostat (a large type of continuously variable resistor still manufactured but not now in common use) thereby obtaining a continuously variable voltage. In practice the unit was not so different from the device used to run electrical arc illuminators, but had the added benefit of using lower voltage at the output (and using a bulb rather than a cumbersome carbon rod gap).

Using an autotransformer with a B&L Research Illuminator means I don’t have to spring for a supply that runs into the hundreds of dollars even when it does turn up for sale. It additionally means not having to worry about setting fire to the workbench, antique electrical apparatus isn’t known for its safety. Furthermore, autotransformers are always constructed with a fuse, which means that in place of the standard 20a slow-blow fuse (as a rule fuse amperage is identical to the rated current) I can use a fast-blow fuse rated for the amps drawn by the lamp being employed, and add a further level of protection for my bulbs filament.

Beyond that there’s the convince factor. The autotransformer is able to supply the required power for every illuminator I have, everything from the 120v Optilume, to the 115v lamp in the Spherical Illuminator, or down to the 6v halogen in the BalPlan. Even the high intensity 6v 18a (think about that, 18a, the breakers in your utility room are probably only rated for 15a on a lighting circuit!) bulb in the the Research Illuminator. So should we throw out the power supplies we do have in favor of an autotransformer? Of course not, but we should be mindful of it as a safe an effective option for feeding power to a microscope lamp of a variety of illumination systems.

Next time something with pictures, I promise. -K

Light Source Power Supply Anomalies

I’ve been meaning to write about power supplies for some time and a recent exchange reminded me of one of the reasons I was initially prompted to. Anyone who’s frequented this odd little website is aware of my feelings concerning used microscopes; in the words of a breakfast cereal mascot “they’re great!” One thing that is perhaps not so great, completeness. By this I of course refer to the tendency of second hand stands to be somewhat incomplete, particularly as regards light sources and power supplies.

Now the absence of a lamp housing and mount should, as a rule, be considered a deal breaker for a stand that requires one. Very rarely, one might find and recognize a needed lamp housing but the search is liable to be complicated by sellers who are uninformed and so list the item under difficult terms or worse by informed sellers who know the proper terms and therefore the rarity and value of the item. This isn’t about lamp housings though, this is about another component that if missing does not disqualify an otherwise complete or desirable stand from consideration; I write of course of the lamps power supply.

Another enthusiast contacted me with a question about a B&L transformer. As I looked through the manuals for a part number I noticed something interesting. Two versions of the manual for the Dynoptic & DynaZoom had two different sets of published voltages! One version of the manual described the five taps as having the first set of voltages, the other the second. Oddly enough each manual recommended the same GE-1634 lamp.

  1. 1v – 2.2v – 4.5v – 9v -21v
  2. 12v – 14v – 16.5v – 20v – 25v

Admittedly, that’s a 20v bulb, so it’s entirely possible that two version of the transformer were made. Somewhat more unusual is the lack of a specific part number listed in either manual for the transformer itself. B&L at one time or another assigned part numbers to everything from screws to shims, so I’m a little concerned that I only failed in my search because I didn’t read as carefully as I might have. I took a multimeter to each of the corresponding transformers in my collection and both proved to me of the higher voltage varieties. It’s not at all uncommon for a microscope illuminator to provide higher voltage than that for which the bulb is rated. In older textbooks and even on some modern transformers the final tap, or range on continuously variable transformers, is marked as “OV” for over voltage generally called over run. The fact that the second set is so much higher than the first would tend to disqualify the first transformer for photomicrographic work. I’d go so far as to say the binocular heads should not used with the first transformer if one intends to use a daylight filter, and the second shouldn’t be used for visual work without one, or at least a set of neutral density filters.

What I’d like to point out, is not that the published voltages of a transformer may not line up with a transformer that “looks like” the one in hand, or that is available for purchase. Rather, that the important thing is the supply provided by the transformer and its suitability not only for the bulb employed but also the intended use. It’s a simple thing really, and something that might be forgiven for someone who’s only had to deal with common lightbulbs of the sort had at the average home store or hardware. Where then should the enterprising microscopist begin in outfitting a microscopes illumination system? With the correct bulb. The correct bulb will be mechanically compatible with the bulb holder and lamp house as well as of the rated wattage.

I write of wattage because at the beginning the most important and most frequently overlooked characteristic of a bulb is the heat which it will put out. Over high wattages will present a fire hazard, apart from the potential damage to a stand one might also damage the eyes, so do consider the wattage when choosing a replacement bulb. If at all possible always use the bulb recommended by the manufacturer, or a mechanically compatible bulb of lower wattage.

And this weekend, the part I’ve been meaning to write! -K

Wait, is that… blood?!

Just buy a used sofa off a guy with a van on craigslist? Find a knife stained red under the floorboards while insulating the attic? Enjoy CSI but hate how the show is super bad about science? Well, let’s play a game I’ll call: IS! THAT! BLOOD?

A little while back, 1853 to be specific, a fellow named Ludwik Karol Teichmann devised an amazingly simple and accurate test for blood. It’s called the Teichmann test or sometimes the hemin crystal test. It’s pretty simple to identify fresh blood from say, stage blood (corn syrup and red dye) or ketchup; blood has red cells and white cells, just make a smear and take a look under the microscope. Old blood, from as little a a few minutes to a few hours can be much harder to identify.

A great many things have cells and are red or brown when dry. Meaning simply re-hydrating a stain and checking for cells under a microscope is not enough to identify blood. If we were on television we could shine a blue light on it or poke it with a stick to test for DNA. Fortunately, human blood contains a number of compounds that are unique to it, and human DNA is just one. Hemoglobin is another and the Teichmann test, acts upon that hemoglobin to form easily identifiable hemin crystals and it works on even decades old samples. Most folks have everything they need right in their kitchen. If you don’t, a five dollar bill is enough to collect the needed supplies from the nearest grocery.

The Materials

Acetic acid, anything from the purest laboratory grade glacial acetic acid to plain old white vinegar will work.

Sodium chloride, iodized salt will do but if there’s a choice it’s better to go with pure sodium chloride-pickling salt or Kosher salt for example.

Something that may or may not be blood.

A glass slip and cover.

A heat source-anything from a candle to a Bunsen burner will work.

The Process

Take the sodium chloride and pulverize it. Only a few grains of salt are needed and the smaller it can be crushed the easier things will go later. For this reason the flake style Kosher salt sold in the baking aisle can be. the best choice. Table salt grains can be effectively pulverized between two slips pressed together and rubbed gently over a third.

With a scalpel or razor knife take up a little of the material to be tested and scrape it onto the slip with the pulverized salt. Only a very little material needs to be used and the proportions of the salt to the possible blood are not important.

Using an eye-dropper introduce a few drops of acetic acid and place a coverslip over. Heat the slip gently just until bubbles are seen to form. It is not necessary or desirable to boil the solution. If the area under the cover should begin to dry out from overzealous heating a few additional drops of acetic acid may be introduced without issue.

Observe the slide under low power-a 10x objective and ocular will do.

The Result

Hemin crystals are elongated hexagonal crystals which will appear ruby colored under daylight corrected illumination. If the salt was not perfectly dissolved in the acetic acid the hemin crystals will be seen to form closely around the un-dissolved salt crystals. This makes for unattractive photomicrographs but has no negative impact on the results of the test. Let’s take a look:


Hemin crystals formed in a positive result for the Teichmann test

So it would seem: YES! IT’S BLOOD!

What now?

If someone was going to aspire at being a sleuth they could prepare in advance a saturated solution of sodium chloride in acetic acid and cary it in a dropper bottle. To do so place 10ml or more of acetic acid in a test tube and introduce sodium chloride a few grains at a time while heating the test tube over a burner. Continue until a small amount of sediment is built up at the bottom of the tube. Allow the solution to cool and place into a stoppered dropper bottle.

In the field a scraping of the material to be tested is placed on a slip and a few drops from the bottle introduced to cover. Place a coverslip over the whole. One will still need to heat the solution to induce the formation of hemin crystals but something as pedestrian as a cigarette lighter will do the job. Observed with even an inexpensive field microscope the results will be obvious.

Sectioning part: II freehand

Although a double edged razor of the type sold for shaving is best for free-hand sectioning, owing to its more stringent manufacturing controls and increased thinness, for initial attempts a single edge utility razor blade is recommended. The thicker blade and single edge contribute to a sense of safety on the part of the nervous practitioner. In either case, the process is the same. One should first toss out any thought of force or slow pace.

For this example a pine needle is firmly pressed against a glass slip with the nearly vertical thumb of one hand. The glass slip is oriented on a slight angle, such that it is in line with the arm which is owner of thumb holding the pine needle against the slide. The reason for this orientation is that it permits the other arm, and the hand which will be holding the razor is easily brought at a right angle to the other. The position of the arms being important as it is the shoulder and elbow of the blade holding arm that will be moved to make the cut. Using the long bones of the arm as something of a pendulum contributes to the smoothness of a cut section and uniformity of thickness. A chopping cut, or wrist controlled section is sure to be too thick, or horribly distorted.

With a drop of water or two introduced to the slip the needle is first trimmed to expose a cut surface. The blade is then placed so that its flat surface is against the thumbnail of the opposite hand. As the arm holding the blade is drawn backwards the blade is allowed to cut a section. Although the blade should be held firmly force should not be applied to make the cut, allow the shard edge to do the work. Any likely sections may be left of the slip or transferred to a second slip for inspection on the microscope. In either case one should not allow the sections to dry out.

I have never acquired the skill for making a good free hand section, but have managed workable results when making cursory examinations of objects I’ll later section with my B&L sledge style microtome. Here are a few images of comparatively “good” results.

One can see that I managed a reasonable thin section but failed as regards uniformity. This is not clear by examining the sections with the naked eye but is glaringly obvious on the microscope. The probable reason for this is down to my not holding the razor perfectly vertical. I’m satisfied with the work of two minutes as compared to the days labor of processing a specimen for the microtome.

Sectioning part: I Theory

It’s bound to happen sooner or later, particularly in a classroom setting where the microscope is not a chosen pursuit, one will run out of things to look at. There’s only so many things that naturally exist in a form that’s suitable for observation with the compound light microscope. Newsprint, onion skin, insect wings, pollen, pond water, and blood, are enough to occupy the interested for a lifetime while others are sure to tire much sooner. Lucky for the instructor, most pupils can be trusted with a knife.


Principles of Sectioning

Transmitted light microscopy is all about the specimens capacity to permit the passage of light from the illumination source through itself and onwards into the objective, ocular, and finally the eye. Some materials are able to permit the transmission of light, more or less, in their natural state, onion skin and Elodea leaves are great examples. Onion skin is comparatively easy to separate from the whole in a sheet one layer of cells thick and many types of Elodea form leaves that are only a single layer of cells. Other materials must be mechanically manipulated to form a suitable specimen. Insect exoskeletons may be macerated, pressed, and cleared. Blood may be smeared. Minerals may be ground. Much else in the natural world, most organic materials, may be thinly sliced.

The thinly slicing of materials as a practice is called sectioning and those materials after sectioning are called sections. An ideal section is thin and uniform with a thickness that is matched to the depth of field of the objective which is used to observe it. In practice a section is apt to be rather thicker than the depth of field but this is generally an acceptable defect provided that the section is uniform and as thin as possible. One may set themselves up for success in regards to uniformity by beginning in sectioning with small objects—a pine needle rather than a branch, the thin tip of a carrot rather than the thick root.

The thinner the section the better the resolution of the resulting visual image. When one focuses on the section the microscope may only focus on the materials that exist within the depth of field of the particular objective, all else exists outside the narrow range of the objective and obscures the image—one will not see the fuzzy and out of focus layers, they will merely lower the sharpness of the image.

Practice of Sectioning

In a professional setting specialized, and nowadays fully automated, devices take care of sectioning; a sample goes in one end as a complete object and comes out the other as a finished slide. Manual apparatus for the preparation of sections are called microtomes (micro for small and tome from the Greek through Latin and French for section) which hold the material and assist the operator in obtaining the thinest and most uniform sections possible.

Microtomes are more often than not used with specimens which have previously undergone a series of preparatory treatments. The object to be sectioned is first dissected from the whole into a manageable portion. It’s then dehydrated and fixed in alcohol to completely remove all moisture and stop all biological processes of the cells. From here the alcohol is displaced by a solvent of paraffin wax which is in turn displaced by the paraffin itself. The paraffin (or other medium) acts to enclose and infiltrate the specimen supporting both the internal and external structures of the object. Properly infiltrated specimens are preserved perfectly and may be stored indefinitely—tissue samples treated by this means have been used successfully in genetic paternity testing decades after preparation and centuries later in the case of infectious disease research!

With their cytoplasm replaced by a supportive and preservative media infiltrated specimens are placed into a microtome and sectioned with exceptionally sharp blades called microtome knives. Some microtomes make use of what’s called a chisel microtome knife which is in essence a large, wedge shaped, razor blade. In some types of microtome the blade is held in ones hand, or by the microtome and moved against the specimen. In other types of microtome the blade is stationary and the specimen is moved against the knife. The microtome knife may also take the form of a cut-throat razor (although the blades have a specific cross section that differs from a shaving razor) or even a disposable razor blade.

The individual wishing to section specimens at home need not obtain a complex and expensive microtome or exotic knife, a simple disposable razor blade and a steady hand is all that’s required. The specimen to be sectioned is held securely in place in one hand over a glass slip. A few drops of water are introduced to the slip and one then slices the specimen with a smooth and speedy motion of the razor. The water serves to maintain the integrity of the sectioned portions which would otherwise dry out in an instant and the quick motion of the blade contributes to minimal distortion.

The razors that work best for sectioning are the thin and precise type sold for use with a safety razor—look for them in the shaving aisle of the grocery. Because razors of that type have a blade on both long surfaces one is forced to hold them delicately and is not able to apply excess force with sectioning. Far from making the activity more dangerous this contributes to safety by ensuring that one is not sectioning tough and obstinate materials that will lead one to slip and slice a finger. After a few sections are made, a likely one is transferred in a drop of water to a second slip and a coverslip is introduced.


Large Format Photomicrography part: XIII

If someone was of a mind to get into large format photomicrography, I’d say go for it. I knew, and still do know, next to nothing about photography. I can do it. Sure, I’ve got a leg up in that I have some gear that does most of the work but even still, a cardboard box will do in the end. Just let it be an adventure.

What to Buy and Where

f you’re going to be getting a number of items shop around. If you buy several things, try and get them from the same source, in the end you might spend a dollar more here or there but you may well end up saving on shipping. If you’re looking for accessories, buy used. A used 4×5 film holder is always cheaper used than new, often less than $10.00 including shipping. For consumables like film, be willing to buy more than you need. A box of 50 sheets of 4×5 works out to $0.70 a sheet while a box of 25 usually ends up at over a dollar a sheet.

4×5 film: Get the cheap stuff. Don’t worry about buying the high speed stuff either, 100 ISO is fine. Amazon works but check out specialist suppliers like Freestyle Photo and B&H.

Paper: Foma. Get the stuff from wherever but buy Foma. Foma is a Czech company that makes all manner of photographic materials. They also happen to be widely available and one of the few companies that still makes a paper slow enough for contact printing on one of those old contact printers.

Chemistry: Pick a developer and know that you will form an unhealthy loyalty to that developer. I picked D76 and love it, I feel like it’s simple enough I will take a shot at making it myself. Can I use it on paper? Yes. Color film? Yes. Expired film? Yes. It’s what works for me. Whatever developer you pick will work for you, don’t stress about it. Same goes for the fixer. All developers develop, all fixers fix.  Pick up a bottle or a pack from Amazon or one of the retailers mentioned above. Then check and see if it isn’t available cheaper on Photographers Formulary.

Changing bag: Buy new. Used ones might leak light and depending on what they’re made of might be devitrifying on the inside. Just buy a new one and use it till you die. It’s going to last forever.

Daylight Tube/Tank: Buy used. My recommendation is the Color By Beseler #8912. It’s great it can do 4 sheets of 4×5, 2 5×7, or an 8×10 and only drinks an ounce and a half of chemistry a shot. If you think you won’t get the bug, and will only do one at a time get an Ilford Cibachrome daylight print tank, smaller is better but get the one that will handle the biggest size you think you’ll need.

What to Read

Anything by Steve Anchell. The Film Developing Cookbook and The Darkroom Cookbook are excellent. You may be surprised that even if you buy used neither of these books are really available for less than $20.00 and that is not at all overpriced. They’re that good. They aren’t something you’ll want to sit down and read cover to cover but they are something you’ll want to refer to again and again.

Black and White Photography by Henry Horeinstein. But you want to shoot in color? Get Horeinstein’s book. Just do it. Light is light and while color is a bit different Black and White Photography is the sort of firm soil I wish I had started out with.

Apart from that there’s tons of stuff out there on the web but it tends to be… well, there’s plenty of great stuff out there. There’s also plenty of absolute trash. Folks who will actively oppose the efforts of anyone who doesn’t pursue photography with the same goals and intents as they do love the internet. I wonder how much of that attitude is the dependent of those people who fought tooth and nail in the early days of photography to give it the imprimatur of art and how much is just the effect of the internet on otherwise lovely folks. Never mind, the less said about insufferable people the better.

Try the people on Flickr, really most folks are nice and the people who chose to associate with the Film Photography Podcast in particular are wonderful. Try the community of people at Lomography as well. There’s something in the attitude of both groups that is so positive and infectious!

In Conclusion

I hope someone out there who never thought they could try large format photomicrography, or who was despairing at only having a low resolution eyepiece camera see’s just what is possible. If you have the gear you can do it. If you have the desire and a shoebox you can do it. Dress up your science fair project, enrage your photography teacher, make a unique gift, or an intriguing photo for the mantle. Film photography is now truly a part of my microscopy and a part of my life. As long as film in being made, I’ll be shooting it.

Next time, freehand sectioning! -K

Large Format Photomicrography part: XII

One thing I’ve learned on this little project is that I’m certainly no photographer! Most of what I know of photography has been gleaned from a handful of books and a few dimly remembered classes at school so I hope anyone who is enough of photographer to call themselves one will take this next bit in the spirit in which it is intended. Photographers are crazy!


Photography is amazing, it’s a powerful, scientific medium and profound means of artistic impression. In the right hands a camera can be a paintbrush or a chisel and mallet. A camera can shout with the report of shotgun or whisper as the falling of snow. It’s not lightly that I say some of the efforts of photographers just make no sense to me, I’m referring to, of course, macrophotography. Some of the accoutrements of macrophotography make sense to me. Specialized lenses that are specifically constructed for high magnification and short working distance are perfectly reasonable in my mind. Even those macro extenders and bellows units that extend the distance from the rear element to the film plane and thereby reduce the minimum focusing distance make sense to me.

I can’t even begin to understand the motivation for some other macro accessories. There are society thread (microscope objective) to camera body adapters that I have to call absolutely insane. This is asking for a bad experience and all but ensuring the objective will be badly damaged. Then there’s so-called lens reversing adapters, or macro-reversing-rings that seam to me with my meager photographic knowledge to be an equally dangerous abuse of a camera lens. Maybe these sorts of devices are sold more for novelty than anything else, or perhaps with my microscope slide orientated vision I have trouble seeing how these are best used.

To my mind if a photographer wanted to have a crack at photographic a microscope slide they’d need a box full of macro optics and a finely controllable vertical tripod of some sort. It seems far easier to my mind to just use a microscope. No, I don’t mean a forty pound monster like the BalPlan or any expensive trinocular microscope at all. I mean a flea-market bargain or a school surplus, monocular, two objective microscope.

Large Format Small Budget

A photographer probably has access to a cut film holder and apart from that all any aspiring large format photomicrographer needs in a microscope. Everything else (not necessarily including the slide one wishes to photograph) is likely already on hand. Today, I used an Amazon shipping box and a desk lamp to get a large format negative of a pine needle section. I supported the box that acted as my camera on a ring stand but I could have used a couple stacks of books just as well, it may have even been more stable if I had!

I took my little shipping box and tapped it up with some strips of duct tape. Next, I traced the outline of my film holder on one end of the box. That done, I measured out a rectangle just under 4×5 inches which I cut out with a razor. On the opposite end of the box I cut a small hole in the center. The hole I cut in the center was only just the size of the ocular, I wanted to keep the hole small enough that I didn’t need to bother with any sort of baffle or light-seal around the eyepiece. I should have used a box a bit longer, a shoe box would have been much closer to the ideal. A sufficiently long box would provide enough extension as to be par focal with the virtual image seen by the eye at the eyepiece. Additionally it would have been a good idea to use a projection or photographic eyepiece so the real image… well lets keep it simple.

Instead of some fancy ground glass a sheet of lean with the protective cover was used but I might just as easily have used a sheet of wax paper. I didn’t bother with any light proofing or consideration for stability. A layer of fleece glued to the face of the box where the film holder would sit would be a useful improvement as would a layer of black paint on the inside of the box—good ideas if I ever find a longer box to use.

The Process

  1. Load a sheet of film into the film holder in a changing bag.
  2. Position a desk lamp with a frosted bulb on the table with the shade oriented to direct all the light downward, place it about 8 inches from the mirror of the microscope.
  3. With your eye at the eyepiece manipulate the mirror so that the light fills the field of view.
  4. Place a slide on the stage and bring the microscope to visual focus.
  5. Place the “camera” over the eyepiece and stabilize it with a bit of masking tape.
  6. Place the focusing screen (wax paper, ground glass, etc.) over the large opening.
  7. Focus the microscope such that the image on the screen is clear.
  8. Turn off the room lights (the darker the room the better, but just enough light to see is ideal).
  9. Remove the focusing screen and turn of the desk lamp without moving the lamp or “camera”.
  10. Place the loaded cut film holder over the opening of the camera and gently pull out the dark slide.
  11. Briefly turn on the lamp to make the exposure and then quickly turn it back off (exposure times up to 5 seconds are reasonable for 100 ISO film, a 40 watt bulb, and a low power objective).
  12. Carefully replace the dark slide.
  13. Process the film.

The Result

I put this whole thing together in about 15 minutes. It took me longer than that to develop the film! Here’s the scanned negative together with an edited inversion in lieu of a print.

Not bad considering I didn’t make overmuch of an attempt at being precise. I should have used a longer box or a more powerful eyepiece, something to raise the magnification enough to fill more of the frame with the specimen. I might have used a shorter box if I enjoyed the look of a circular vignette. A shorter exposure would have been good as well, the negative is very dense and the only thing that saved it was the very dark stain in the specimen. With the 40 watt bulb, 10x objective, and 5x ocular I’d wager a 2-3 second exposure would have been closer to an ideal result. Still, not bad for a microscope that I picked up for under $20.00 USD.

Next time, the wrap up! -K

Large Format Photomicrography part: XI

At this point I sort of know what I’m doing as far as large format photomicrography goes. Which is to say I can put a slide on the stage and reasonably expect to end up with a serviceable print. For anyone who’s been here through the entire series to this point it likely feels as if this has been going on forever. All told what with the demands of work, other interests, and responsibilities, on any given day when I picked up a film holder I’ve probably spent no more than an hour on the project. Between reading up on things, photographic work, operating the scanner, and making notes the whole things been rather a rush.

Forgive me then if I step back a minute and put a couple scribbles up on the ‘fridge.

Two Scanned Negatives

Two Scanned Prints

Pretending I’m A Photographer

The negative of the lilium ovary section above is a bit thin but has enough density to provide all the detail that’s present in the visual. It was a 1/15th second exposure which I was a touch concerned would be a bit too long with the lightly stained section. The negative of the zea stem, 1/8th of a second, is about right but has a defect where something (I checked later and it was a mote on the System II relay lens) obscured a portion of the negative.

I made up for the lower density on the lilium negative with a bit of a longer exposure on the contact print, I might have over-done it a bit but I’m not unhappy. With the significantly more dense negative of the zea I used what I felt would be a long enough exposure for the contact print, just under two minutes. If I had the presence of mind to I might have dodged the mote while I made the contact print. I expect I’ll give that a try if I ever make a second print from that negative.

With photomicrographs large format really opens up the possibilities for the microscopist. In this the day of digital cameras and desktop photo manipulation one can capture an image with an extensive depth of field and an enormous field of view. Their isn’t really a way to expand the depth of field for the chemical photographer short of better objectives. The field of view can be greatly expanded by making the switch from the classic—notably called miniature historically—35mm format to a medium format 120 film, or low end large format like 4×5. From there the step up to 8×10 would mean capturing the zea stem with a 20x objective or the entire lilium ovary with a 10x. Considering that it’s somewhat strange that in the large format photomicrography did not last quite so long as 35mm, which oddly enough still has a presence in electron microscopy to this day.

Up to this point I’ve made use of standard equipment. Earlier I theorized in an off hand way about how one could knock together a 4×5 camera for a basic monocular microscope without too much trouble. For my next trick, I’ll give that a shot! The target audience would be someone who happens to have a microscope and a friend who shoots 4×5, or someone who shoots 4×5 and wants to give extreme macro photography (photomicrography) as go. I’ll skip over the business of developing the negative, as that grounds been covered, and focus on seeing if I can get a negative at all with a shoebox and a few odds and ends.

Large Format Photomicrography part: X

For absolutely silly reasons I’ve done this a bit out of order. By all rights I should have exposed and processed my first contact print in my improvised darkroom using open trays. Under the light of my spray painted night light “safe light” I could easily set up my exposure and observe the level of development as it progresses. That would give me a ready idea of the required development time and let me somewhat adjust for over or under exposure by pushing (extending) or pulling (limiting) the developing time.

The Changing Bag Contact Prints

Loading everything into the changing bag wasn’t too terrible. The worst part of the whole thing was being entirely unable to see if I was aligning the contact paper up with the negative. I had to resort to tracing the outside edge of one of the metal slats and slowly bringing the edge of the paper up to it. It was all the more hard as the changing bag prevented me from fully opening the lid of the contact printer. The lid fails to stay in the open position unless fully open so it was all the more difficult as a result. Patience was the key and below is the first result.

1 sec 2x40

Overexposed contact print

This first contact print (at left) was made using the decades old dual 40 watt Mazda bulbs that were installed in the contact printer when I received it. The exposure was made for one second after which it was processed in Dektol for just under 45 seconds with constant rotary agitation followed by a two minute plain water stop bath. The print was then fixed for two minutes using a 1:7 dilution of Kodafix. As may be seen the print is exceedingly overexposed having hardly any definite texture in the legs of the opilione (daddy long legs spider) and none in the body. I drastically underestimated the brightness of the lamps serving as the light source. I took into account that with the contact printer the distance from the light source was easily ten times shorter than that one would use with an enlarger and working the manufacturers data sheet describing the paper as 30 times slower than normal papers I selected a one second exposure and well, at least it was educational.

3 sec 2x15 d

Unevenly exposed contact print

With the above as a reference point I looked at what I had available in the way of medium base (medium Edison screw, or E27) bulbs. Failing to find anything less than 40 watts with a frosted or opal glass I settled on a pair of 15 watt night light sized clear bulbs—the type used in the B&L Opti-Lume illuminator. Apart from being significantly lower in wattage, the night light bulbs are much smaller physically and have a shorter total filament length. In the second print (at right) I used an exposure time of three seconds and processed the print as above. The results were better than those had in the first contact print but still quite a bit off from acceptable. The mounting hardware in the contact printer is on one side only so that two full sized bulbs have their filaments centered beneath the frosted glass. The smaller bulbs were far from centered and an internal wire partially occluded one of the bulbs.

Tray Processed Contact Prints

3 sec 2x15

Preferentially developed right corners with tray development

Resolved to get something much more like an acceptable print from the contact printer I adjusted the position of the internal wire and repeated the three second exposure with the night light bulbs and a different negative. This time I worked in my improvised darkroom and processed using Dektol and Kodafix in an open tray. I used a plain water stop in a third larger tray. Trying to be too clever for my own good in my first attempt at tray processing I sought to overcome the effect of the off center bulbs in my contact printer. I used roughly a minute and a half of total processing time and for nearly a third of that I held the side that corresponded with the bulbs out of the tray using my tongs and preferentially developed the opposite side of the print. The results as seen at left aren’t more even as a result, if anything they’re less. Rather than being more even the one side is significantly less developed overall and there’s somewhat less overall contrast. With little experience on the matter I’ll tentatively attribute this to the far lower rate of agitation I was able to achieve in the trays as compared to the constant agitation in the rotating print drum.

At this point I decided to make an attempt with my photographic enlarger. I had actually bought the enlarger on whim on the off chance that I’d one day be sufficiently enthusiastic to have a go at putting together a darkroom. The portable enlarger by Ilford isn’t able to handle a 4×5 negative for enlarging but it will work for a contact print. I began with a 15 second exposure time and processed in trays. When after the first minute of developing nothing was visibly happening with the print I started to think I must have had the print upside down in the printing frame. I tossed it in the general direction of the water tray and moved on, setting up another sheet of contact paper on my printing frame. Then I noticed something on the print that lay in the sink beside the water bath, it had developed to a limited extent! I gave it another couple minutes in the developer and started to see it a bit more clearly, at which point I put it in the water stop bath and thence into the fix. The result is below on the left. I left it in the water bath while I exposed the next attempt.


For the second print with the enlarger I used an exposure of 30 seconds, two minutes in the developer, two in the water stop, two more in the fix and then into the water bath. I ran the water from the sink into the water tray while I poured the chemistry from the trays back into their storage bottles. I made small hash marks on the masking tape labels of the bottles so that I could gauge the remaining capacity of the solution in the bottles. With that done I took the prints one by one and hung them to dry.