Veils in the Water: “Schools” of Protozoa

“Veil” in week-old sample from Washington Park Arboretum

Sample some pond water, bring it home and let it sit quietly for a week under a light, and you may experience a fascinating sight: ghostly veils and arcs hovering in the water.

After sitting quietly under a desk lamp for a week, a little thrift store aquarium full of water, pond plants, and algae from the Washington Park Arboretum developed these eerie white arcs hovering above a clump of algae (See arrow). They are very difficult to record -it is frustrating to persuade the autofocus on a camera to lock onto them – but with multiple exposures and much processing, it was possible to create a credible image.

Carefully inserting a pipette, I sucked up the area, capturing a few milliliters of what appeared to be clear, completely unremarkable water – but again the microscope showed another world. Spread across the field were a swarm of tiny green, peanut-shaped organisms, each swimming with a peculiar curliqued movement: several rounds in a circular spiral, then a rest, then another circle, then a pause, and so on:

Still images revealed small, peanut-shaped flagellates, each with two curled flagella:



These are tentatively identified as Cryptomonad algae, possibly Chilomonas (thanks to Wim von Egmond and Vincente Meneu, Amateur Microscopy Facebook group).

I have noted two other occurrences of protozoan schooling.  In one case, delicate but very distinct cloudy veils formed in a mini-aquarium of pond water after it had been set up for several weeks; unfortunately, I was not able to take a sample.  However, in my very first observation of protozoan veils, a 3 cm, cloudy band appeared one morning against the glass of my original mini-aquarium.  A 4X microscope objective on my Samsung cell phone camera showed a school of tiny organisms:

Protozoan school in aquarium, 8-10X

Under the microscope, the tiny dots resolved into a swarm of dinoflagellates:

 Only one other microscopist has commented on this phenomenon on the web:

“In 2006 I noticed a discretely-shaped “veil” about 2 – 3 millimeters in extent, moving through the water of a mini-aquarium where I was also looking for hydras. The shape formed was like a section of an auger bit, or the twisted form of a DNA molecule (but only about 1/2 to 3/4 of a “twist”), and sometimes like a “saddle shape.” It also turned out to comprise something that looked most like Chlamydomonas, but I did not have good enough lighting (mirror understage only) to see bacteria.

I have often wondered since then, how many people are aware of protozoans swimming in organized schools, and what mechanism they use to communicate their positions to each other. The shapes they formed had VERY definite edges (L. Hizer, personal communication, Facebook comment, 2016/09/24)”.

As noted by Mr. Hizer, this phenomenon raises fascinating questions, .  How are these swarms created? Is there a mechanism by which the cohesion of these protozoan “schools” is maintained?  If so, on what basis does it function?  Chemotaxis, light sensitivity, or some unknown mechanism by which protozoa sense each other, rather like flocking behavior in  birds?

The slide of the Cryptomonads also showed many slender bacteria – could this swarm be a collection of organisms clustering around a food source, rather like killer whales around a school of herring, or a response to some bacterial metabolic product?

Many bacteria in a Cryptomonad swarm.  What is their significance?

The most likely answer comes from research on something called “microscale nutrient patches“.  Early modeling studies on movements of plankton, especially unicellular organisms and marine bacteria, considered lake or sea water as a homogeneous medium, but modern thinking suggests that nothing could be further from the truth.  Consider that our environment – the air around us – is anything but homogeneous.  If you ride a motorcycle outside the enclosed environment of a car, you will experience dozens of micro-environments in the course of an hour’s ride:  hot and fragrant near a newly-mown hayfield, damp and cool in a gully, smelly behind the exhaust of a bus, windy and salty near the ocean. Not to mention the aroma wafting downwind from a roadside rib joint…

Now transpose this image to the Bering Sea or a pond basking in the sun.  Not only is the chemical environment different at different depths, but a large body of recent research suggests that every plant, invertebrate, or single-celled organism releases a cloud of chemicals in its immediate environment (see Blackburn). Many of these are vacuumed up by motile bacteria  as food, while others serve as attractants for other unicellular creatures.  Observing the motile bacterium P. haloplanktis, Vahora noted chemotaxtic attraction occurring within seconds  when a single cell of the alga Thalassiosira was introduced into the culture.

On consideration, this is not surprising; the world is full of gradients of chemical attractants and repulsants.  Lemon sharks can detect tuna oil several hundreds of meters away at a concentration of one part per 25 million.  Feeding releases blood and body fluids that attract more sharks, etc.  Anyone who has observed a family hovering outside the kitchen while the Thanksgiving turkey cooks should immediately grasp the concepts of chemotaxis and nutrient patches.  Coco Chanel has made millions from the principle of chemical attraction.

This phenomenon has been extensively studied, and has been documented in many species. The unicellular flagellate algae Chlamydomonas and Dunaliella tertiolecta have been shown to be chemotactic to several substances, including ammonia. The ciliate Paramecium has become the model organism for these studies on protozoa, with development of commercial vat cultures and creation of genetic mutants lacking various chemoreceptors (see the excellent review by Houton and Preston).  Such studies can be of enormous importance, as they may shed light on how cells in general (including our own body cells) sense their environment and react.  Chemotaxis plays a critical role in wound healing and embryonic development, and defective chemotaxis is thought to have significant involvement in asthma, atherosclerosis, and chronic inflammatory diseases (see Kamp et al.)

So how does chemotaxis occur, and how do the dinoflagellates and algae in our specimens congregate to form swarms?  Mechanisms seem to be different between bacteria and protozoa.  Bacteria execute what is called in statistics a “random walk“, where locomotion consists of a “run” and a “tumble.” With E. coli, “runs”, or movement in a straight line, occur when the flagella rotate counter-clockwise, and “tumbles”, or directional changes with no forward movement, occur when the flagella rotate clockwise. “Run” length, speed, and frequency of “tumbles” are all variable.  When a cloud of bacteria finds itself in a concentration gradient of nutrients, these variables change in response to the composition of the immediate environment.  Movement up the nutrient gradient is favored, leading to a “biased random walk”  with the cloud drifting toward the food source:

Bacterial “random walk” up nutrient concentration gradient
Bacterial random walk toward food source (courtesy Schmidt and Kardan)

Bacteria respond to external chemical stimuli through transmembrane “methyl-accepting chemotaxis proteins” or MCPs.  After binding to specific chemicals in the surrounding fluid, MCPs transmit the message across the membrane into the cytoplasm, where Che proteins are activated.  One Che protein type interacts with the “Flagellar Switch Protein”, changing flagellar rotation from counter-clockwise to clockwise and causing a tumble.  Changing tumble frequency biases the random walk toward the higher end of the gradient, and the whole population drifts in this direction.  This is discussed in Wikipedia’s article on chemotaxis.

Bacterial Transmenbrane Methyl-accepting Chemotactic Protein (MCP)

This is how a cloud of bacteria move.  So what happens in an animal or plant, non-bacterial cell, when chemotaxis occurs?  You can thank all those little creepy organisms  in the soil and water billions of years ago for doing the basic R&D. The mechanisms they developed seem to have worked, because we are still using the same chemicals, whether you are a slime mold, an annelid worm, or a Member of Parliament.  We all use “G Protein Coupled Receptors” or GPCRs, a family of proteins that transmit information across the cell membrane.  GPCRs have a charged N-terminal end on the outside of the membrane, seven fatty helices that span the lipid-rich center of the membrane, and a charged, water-soluble C-terminal end inside the cell.

The N-terminus on the membrane surface (cell surface “receptor”) is configured to bind to a specific chemical or hormone; binding causes a subtle structural shift (conformational change) on the outside segment of the GPCR protein.  This shift in the complex, tangled yarn of the protein echoes across the membrane to the segment inside the cell, where GTP, a signaling chemical or “messenger“, is released, unleashing an enormously complex chemical cascade:

Mechanism by which binding of a small chemoattractant to a membrane surface GCPR releases messengers GTP and GDP, with subsequent activation of actin and myosin fibers via complex chemical pathways.  See Kamp et al. for nomenclature and discussion.

In slime molds and human neutrophils, activation of this mechanism causes the whole cell to rumble forward, “…creating actin-rich pseudopods at the front and retracting the back of the cell using myosin filaments…”  This mechanism is detailed in Kamp et al’s just-published and very complex, but excellent, review article.  In pond critters, this mechanism controls the direction and speed with which they swim toward a food source.  The intracelluar architecture of this process is enormously complicated, and our understanding is in its infancy.

Research on chemotaxis and directed locomotion raises more questions than answers once we get above the level of bacteria.  Paramecia seem to display elements of the both the biased random walk and directed swimming. However, the mechanisms by which higher single-celled organisms interact with their environment are multifaceted, and much more complex than those used by bacteria. Protozoa are large enough that a chemical gradient outside the cell may be reflected in a chemical messenger gradient inside the cytoplasm from front to back; the recently-discovered complex phospholipid PIP3 may be involved in this process.  In addition to releasing internal messenger chemicals, binding of chemoattractants to surface GCPRs causes depolarization and changes in the transmembrane voltage that control swimming (Valentine et al.). Receptors are probably distributed in specific areas over the cell membrane, and there is evidence that they can be synthesized quickly in response to stimuli.

Cilia themselves are covered by specialized extrusions of the membrane, act as sensory organs, and have complex internal functions that involve molecular transport between the tip and base:  “…cilia and flagella are micromachines and they act as cybernetic devices to receive, process and communicate information… (see Linck).  Voltage changes in the ciliary membrane open calcium channels that affect the strength of the ciliary wave, altering direction and speed.  Control of ciliary and flagellar beating, and hence direction and speed of swimming, appears to involve a complex and poorly-understood relationship between ciliary membrane voltage, calcium and magnesium ion movement, and intracellular released cyclicic nucleotides (cAMP and cGMP) (see Andrivon).  Cilia may also be in part coordinated by a system of intracellular fibers and tubules.

These mechanisms have implications for more than the wanderings of protozoa.  A whole spectrum of genetically-coded diseases are now thought to be “ciliopathies“, or inherited defects in ciliary-associated functions.  The body of research in all of these areas is vast and fascinating.

Complexity of ciliary architecture and transport mechanisms (see Emmett et al.)

Amazing that we can couple a little shadowy swarm in a cloudy bottle of water to such profound mechanisms at the root of our own existence!

So what does all this tell us about our little watery veils of algae and dinoflagellates?  The occurrence of these miniature swarms of protozoa is probably associated with a tiny, invisible patch of food.  Chilomonas is a cryptomonad flagellate that has no pigment in its chloroplast (leucoplast) and must survive on soluble compounds.  Its favorite carbon source is acetate, but it will survive equally well on succinate, and also gains nutrition from straight-chain fatty acids and alcohols (see Nisbet, p. 166).  Both succinate and acetate are, however, produced under anaerobic (low oxygen) conditions, and would more likely to be found in sediments and ooze. Another attractant is more likely to be responsible.

It is, however, interesting that the Chilomonas swarm was associated with a proliferation of bacteria.  Were the Chilomonas slurping up waste products of the bacteria, or was it the other way around?  With the other swarm, Dinoflagellates have enormously broad feeding behavior, ranging from photosynthetic species to predatory varieties, so understanding their swarming responses is more uncertain.  Clearly, they were attracted by something in that small area, even though we could not see it.

Protozoan schooling may be much more common than previously recognized.  Even when present, protozoan clouds are very difficult to see with the naked eye.  Like ribbons of smoke rising in the air, these delicate aquatic veils are apparent only with appropriate lighting, ideally from the side.  They would be virtually invisible through the water’s surface in ordinary sunlight.  Furthermore, they would form only in very still waters and be dispersed by even a gentle current.  You are encouraged to look for these delicate structures next time you leave a bottle of pond water on the window sill.


This is the beauty of natural science:  a simple observation of  phenomena in a bottle of muddy water opens a path to principles critical to the function of life itself.

Discovering this delicate and fleeting phenomenon is a testament to the potential value of work done by dedicated amateur microscopists with careful observational skills and a desire to communicate their findings.  Even modest equipment can allow worthwhile observations like these (done with a 45-year old Nikon S-Kt, a cell phone camera, and an international group of helpful fellow enthusiasts).  Professional scientists are too often torn between teaching, grant writing, and gaining tenure, not to mention their own research (often farmed out to graduate students), to have time for interesting natural science.  Research topics are often strongly influenced by what will be funded, and observing pond water is not often the best path up the academic ladder.

The situation is similar in astronomy.  A night of observing time on the Keck Telescope costs $53,700.  Obviously, this is one good reason why new asteroids and comets are often detected by amateurs with basic equipment, time to spend, and no research committees to satisfy.  As an example of the value of amateurs to astronomy, NASA is presently promoting a new program (part of the ORISIS asteroid sampling mission) to engage amateur astronomers in documenting new asteroids, an avenue of research that can now be done with a good 8″-10″ telescope and CCD camera.  “Citizen Science” is well organized within NASA, and there are many established avenues for competent amateurs to make scientific contributions.

Although scientific opportunities outside astronomy or established institutions are miserably infrequent (see Kathlyn Mills article in Wired-UK, and Bruce Bigelow’s still-relevant 1996 article in The Scientist), some formal opportunities exist for citizen scientists. Some of these may provide opportunities for microscopists. is a federal web site sponsored by the General Services Administration and the Wilson Center; as of October 2016, it listed 303  federally-sponsored natural science projects needing citizen scientist participation.  Europe has the European Citizen Science Association, and Australia has the Australian Citizen Science Association.

There is less public attention and less funding for the world of the very small as compared to NASA’s world of the very large, but amateurs are making definite contributions.  Popular Mechanics article, “Inside Amateur Science: The Best in Out-of-Lab Research“includes the work of Ely Silk, who makes fluorescence LED illuminators for microscopists. The Diatom Herbarium of Drexel University and much of the database of New Zealand diatomology were largely built from the collections of amateur diatomologists.  Loren Bahls‘ recent paper details the important role played by “…citizen volunteers…(and)…discusses the process of engaging citizen volunteers in diatom collection and the value of citizen collections in building diatom herbaria, in cataloging diatom biodiversity, and in expanding our knowledge of diatom biogeography, especially in remote regions with difficult access…”

More than anything, there is an enormous need for microscopists who will popularize this essentially unknown avocation.  Microscopy is a better-known pastime in Britain, which has a long tradition in this area.  In the U.S. and Canada, I am met with either fascination or blank looks.  At best, I am viewed as a harmless eccentric; I worst, I am still waiting to be picked up by the police during my midnight marsh wanderings.  There is a great need for people who will set up a chair and a portable microscope under a tree at a lakeside beach and wait for curious children and parents to congregate.  Try it!


American Museum of Natural History.  “Myth 5:  Sharks can Detect a Single Drop of Blood in the Ocean.”

Andrivon C.  “Membrane Control of Ciliary Movement in Ciliates.”  Biol Cell. 63:133-42 (1988).

Blackburn, N., Fenchel T, Mitchell JG.  “Microscale nutrient patches in plankton habitats shown by chemotactic bacteria.” Science 282: 2254-2256

Clark, R.  “Combined Amateur Telescopes for Asteroid Detection.” Polymath Blog, April 12, 2016.

Emmer, B.T., et al.  “Molecular Mechanisms of Protein and Lipid Targeting to Ciliary Membranes.”  J. Cell Sci. 123: 529-536 (2010).

Govorunova, E.G. & Sineshchekov, O.A. “Chemotaxis in the Green Flagellate Alga Chlamydomonas.”  Biochemistry (Moscow) (2005) 70: 717

Haake, A.  “Inside Amateur Science:  The Best in Out-of-Lab Research.” Popular Mechanics Online, June 10,2009.

Houten, J.V. and Preston, R.R.  “Eukaryotic Unicells:  How Useful in Studying Chemoreception?”

Kamp, M.E. et al.  “Function and Regulation of Heterotrimeric G Proteins during Chemotaxis”.  Int J Mol Sci. 2016 Jan; 17(1): 90.  Published online 2016 Jan 14.

Linck, R.W.  “Cilia and Flagella.” In: eLS, Citable Reviews in the Life Sciences, July 2015. John Wiley & Sons Ltd, Chichester. [doi: 10.1002/9780470015902.a0001258.pub3

Nisbet, B. Nutrition and Feeding Strategies in Protozoa. Croom Helm Ltd, London, p. 166 (1984).

Schmidt, A. and Kardan, M. “Chemotaxis:  How a Small Organism finds a Food Source.”

Vahora, N.  “Micro-scale interactions between chemotactic bacteria and algae”.  Thesis, M. Eng., Massachusetts Institute of Technology, Dept. of Civil and Environmental Engineering.

Valentine, M. et al.  “Chemosensory Transduction in Paramecium.” Jpn. J. Protozool. 41:1-7 (2008).

Wikipedia.  “Chemotaxis.”

Wikipedia.  “Lemon Shark”

Veils in the Water: “Schools” of Protozoa

Green Hydras and Simple Lenses: On Being Creative with What You Have

Green Hydras, 15-20X

Imagine that you could eat lunch by just lying out in the sun?  That’s what these green hydras (Hydra virdissima) are doing after moving from their Washington Park Arboretum pond to one of my small desktop aquaria. Green hydras are phototrophic, congregating in areas with highest light to energize their symbiotic green algae.  These little fellows looped over and over up a stem to congregate at the top, which was about 2 cm below the surface under an LED desk lamp.  Then they clustered together like tourists on a beach in Miami, soaking up the rays.

My problem:  How do I photograph this coelenterate tailgate party???  The little stem was about 3 cm from the glass, out of range of microscope lens-based magnifiers and too small to magnify well with a cell phone on macro.  The macro setting on my old Canon point-and-shoot doesn’t go beyond 4X.  I don’t have one of those new digital 10X microscope cameras.  Nor do I have a bellows and sophisticated ultramacro lens for my digital SLR.  So how was I to record this event?

I used a Canon EOS digital SLR (half-frame sensor) with a Tamron 18-50 mm lens (about 35-85 mm 35mm equivalent) at ISO 1600, f/5 at 1/30 second and 50 mm setting. Then I added one of my favorite lenses, the Vivitar Series 1 10X macro close-up lens.

The Vivitar Series 1 lenses are old 35 mm lenses dating from 1975-1995. Vivitar sold aftermarket lenses that they had made by various manufacturers. Consequently, the quality varied from mediocre to superb. A couple, like the 70-210 macro, became legendary and were some of the best lenses of the 20th century.

The 10X closeup lens is a big old single hunk of glass that must be 1 cm thick in the center. Coated and sharp, it has nothing even resembling a flat field, but for non-planar subjects like flowers, insects, and wriggly pond things, it can create some superb images. It was typically sold in a set of four in a nice pouch; these can be found on Amazon or eBay for less than $20.

Using this combination, I was able to do a 10X image from 3cm away through a light film of algae on the glass, then crop to about 15X. I used manual focus at maximum magnification, then took about forty images hand-held on a box through the plane of focus. I didn’t try stacking, as the arms move slowly.

Posting this on Facebook’s Amateur Microscopy site brought up a number of responses, some enthusiastic and some dubious. This prompted the following thoughts on this lens, older lenses in general, and craftsmanship in photography, whether through the scope or out on the side of a mountain.

In today’s world of computer-designed, aspherical lenses and automatic HDR, we can point even a medium-priced camera anywhere and create a high-quality, flat-field image with better shadow detail than I can see with the naked eye. However, in this world where almost any lens is better than the one Ansel Adams used, I think that we have lost a connection with our lenses and their quirks and individual character. And maybe we have lost an element of creativity, too.

I too started work trying to buy the best and sharpest lenses I could find – Zeiss Sonnars and Tessars and Planars. Then I got into vintage cameras and found myself working with much simpler lenses. Every lens had its personality, and I found that I had to know every one like a member of my family. Then, I could make them sing – and in doing so, I stretched my own knowledge and creativity.

The big old Series 1 Vivitar (really a glorified, high-quality magnifying glass) is like that. A very imperfect, non-corrected lens, but also capable of some amazing effects if you just learn to work with its personality. These images show some of its strong points: the distortion creates arty effects in the flowers, and the leaf tips are in sharp focus because they are both in the midfocal plane of the curved region of sharp focus:

These aren’t microscopic images, however, so what does this have to do with microscopy? The first point is, that even though this is a simple and imperfect lens, it provides an easy way to enter that important no-man’s-land between macro and micro. It is an extremely powerful lens that can let my camera work at a distance of 3-4 cm and still take images of tiny creatures like copepods and hydras. Most macro lenses won’t go to 10X even with extension tubes, and working distance is often short with other arrangements such as low-power microscope lenses.

So restrict your subject to what works well with this combination, put the hydras or whatever in the center of the focal field, and think about what to do to make the rest of the picture look good – this is an exercise in creativity.

Secondly, unlike more sophisticated cameras-and-bellows-and supplementary lens arrangements, it’s cheap ($5), rugged, and portable. The more complex arrangements ARE very good, and much more technically sophisticated. However, if the hydras come out to play while you’re in the kitchen, you can run upstairs, slap on this lens, shoot your pictures, and be back before the soup boils over.

I’m living much of my life in hotel rooms, so everything has to be packable and rugged. The Vivitar is not perfect, but where else can you get a gizmo that lets an SLR photograph hydras, costs $5, doesn’t fall apart if you knock it, and can be slipped into your pocket to photograph bees and lichen on the side of a mountain in the rain? But you have to be willing to think creatively about when it will work and when it won’t, and you might just get an unusual image by thinking a step farther about how to make it all work.

But it’s another example of how knowing your equipment and its craft, and really working with what it does and doesn’t do well, can let you do wonderful things with simple stuff. Don’t turn up your nose at a simple device until you really know how to use it.

The CRITICAL point of all of this, especially for new microscopists who may despair of ever making good images or doing serious work, is that microscopy, like astronomy, is a field where worthwhile investigation can be done by serious amateurs with time to spend.  Popular Mechanics published an article on the sophisticated contributions that amateur scientists working outside laboratories are making to modern science; note that one of them is a microscopist  dealing with LED sources for fluorescent microscopes.  Amateurs are making significant contributions to diatom research.

And you may not need the most sophisticated equipment, either.  A Zeiss scope with a full complement of planapochromat objectives is wonderful, especially if you are doing high resolution work.  The ability to do dark field, phase contrast, and other alternative lighting mechanisms is highly desirable.

Join one of the many microscopy societies worldwide


Bahls, L.  “The role of amateurs in modern diatom research.”

Haake, A.  “Inside Amateur Science: The Best in Out-of-Lab Research.”  Popular Mechanics, Jun 10, 2009.  Accessed Sept. 12, 2016.

Green Hydras and Simple Lenses: On Being Creative with What You Have

Skagit Valley: A Natural Hay Infusion

Rawlins Road, Skagit River Delta. Hayfields and roadside drainage ditches.

Exploring the Skagit River delta north of Seattle on the weekend. Found this beautiful spot where hayfields had just been cut. There is a drainage ditch on either side of the road (see image) that was full of hay clippings – looked like it had been there for several days.

Roadside ditch – full of hay clippings, incubating for about a week. A delicious-looking soup!

Sampling the water revealed an amazing proliferation of life: at the least, one can count nematodes, rotifers, various types of ciliates, spirotrichs, dinoflagellates, and the many small green ovoid organisms (Euglena?):

Skagit Valley: A Natural Hay Infusion

Lacrymaria olor and the Litostomata – Beautiful Ciliates with Nasty Dispositions

Images of Lacrymaria, from 1838 drawings by the German microscopist and naturalist Christian Gottfried Ehrenberg, 1795-1876.

31-17-1-Lacrymaria-proteus 38-7-1-Lacrymaria-olor-originall-Trachelocerca-viridis



Lacrymaria is truly a creature out of someone’s nightmares. Little known and underappreciated, this fast-moving, venomous predator extends its neck seven times its body length to engulf its victims, and has no hesitation in taking bites out of unlucky creatures too large to swallow.  The lyrical name of Lacrymaria olor – “Tear of a Swan” – and its slender, willowy profile belie the ferocity of this creature:

Fortunately, this sinuous monster is unlikely to remove a toe the next time you step in the lake – at only 1/10 mm (100 microns) in length, it is one of the smaller ciliates – but it makes up in ferocity what it lacks in size.  Lacrymaria is one of the litostomate ciliates, tiny creatures with unique, highly-specialized mouth structures.

Litostomatal ciliates are fascinating and beautiful, but their habits do n0t make them  good neighbors:

“Litostomatean ciliates are predatory, with many extrusive toxicysts in oral area. They swallow algae, flagellates and ciliates and even rotifers. The cells are very mobile and some (e.g. Lacrymaria) show extreme rapid changes in shape.” (ref.: Rosati et al., J. Eukaryotic Microbiol. 50: 383, 2003).  Courtesy W. van Raamsdonk,

This class is diverse, and includes many  carnivorous protozoa, other, anaerobic ciliates that live commensally in the gut of many species, and the only human ciliated intestinal parasite, Balantidium coli.

Litostomateal ciliates

The exact characteristics of this class have to date been poorly characterized;  Gao notes, “…The class Litostomatea has been traditionally rather poorly defined as having an apically positioned cytostome, uniform somatic ciliation and a non-distinct oral apparatus…”  Unlike many protozoa, where ingestion can occur at any site on the membrane, this group of organisms sports an apical cytostome or “mouth”.  The latter consists of an indentation specialized for phagosytosis, with an underlying support structure composed of two sets of microtubules.  The cytosome is usually, but not always, associated with a cytopharnx:

“…a long, tube-like structure that forms the invagination associated with the cytostome…it is typically directed towards the posterior of the cell, often hooking around a central nucleus. The length of the cytopharynx varies during the cell cycle, however the average length is 8 µm. Much like the cytostome, a set of microtubules form an association with the cytopharynx. Two sets of microtubules follow the path of the cytopharynx in cells. These sets of microtubules form a gutter-like structure that surrounds the cytopharynx…”

“…macromolecules…pass into the lumen of the cytopharynx and are transported to the posterior end of the cell where they are put into budding vesicles that are transported to other parts of the cell. The cytopharynx in this way acts much like a straw that sucks macromolecules to the posterior end of the cell…”  (From Wikipedia, “Cytostome.”

Genetic analysis has upset and reorganized traditional morphological classification of single-celled organisms.  Gao et al., 2016 have recently constructed a classification of the ciliates based on both morphology and several different genetic markers:

Modern classification of the ciliates using combined traditional morphologic criteria and genetic analysis (from Gao et al.)
Modern classification of the ciliates using combined traditional morphologic criteria and genetic analysis (from Gao et al.)

On the basis of modern morphological and genetic evidence, the Litostomatea (on the left in the above diagram) have three subclasses:

  • Haptoria: comprising Lacrymariida, Haptorida, Didiniida, Pleurostomatida and Spathidiida
  • Trichostomatia:  Symbiotic ciliates in the gut of vertebrates. Also includes Balantidium coli, the only known ciliated intestinal parasite of humans.
  • Rhynchostomatia:  The most recently-created subclass, includes Tracheliida and Dileptida.  The distinguishing characteristic of this subclass is the location of the mouth at the base of a long proboscis, the latter covered with cilia and possessing toxicysts used to stun prey.  The Dileptids (e.g., Dileptus) are aggressive predators,  “…equipped with a long, mobile proboscis lined with toxic extrusomes, with which they stun smaller organisms before consuming them…” (Wikipedia, Dileptus).   Dileptus‘ hunting habits have been described as “rapacious.”  The Tracheliida are similar to the Dileptida but vary in the mouth structure and arrangement of cilia.




Anyone who wants to understand this fascinating class of ciliates should not miss reading Vdacny and Foissner’s  “Monograph of the Dileptids (Protista, Ciliophora, Rhynchostomatia)”(  Although focused on the Dileptids, this exhaustive (529 pages), yet very readable treatise is inclusive, well-organized and thoughtfully written.  It proves much information, as well as many drawings and images, of ciliate ultrastructure, that generalize to the structure of all protozoa.


If, as I have, you feel discouraged and ignorant because you struggle to classify tiny creatures neatly into phylum, class, order, family, genus and species, take heart.  Genetic analysis has made thousands of hours of academic study and morphological classification completely irrelevant.  Careful classifications based on overall anatomy, ciliary structure, or the gonadal morphology of worms have been turned on their heads.  See Wikipedia’s article on taxonomy to appreciate the evolution of this science, and the degree of present-day flux in our thinking about the slots into which we put the living world around us.

We have a much improved understanding of the structure of the Tree of Life.  This does not, however, mean that this new information has simplified the life of the microscopist.  All but the most general classification systems are morphing continually as branches die off and new limbs grow.  Those still suffering from an inferiority complex should consult the papers by Gao et al or Zhang et al’s  “Insights into the phylogeny of systematically controversial haptorian ciliates based on multigene analyses” and admire the beautiful, multicolored taxonomic trees and RNA maps.  These would look lovely on a wall or embroidered into seat covers, but tend to induce stomach pain in anyone short of a graduate-level genetic taxonomist.  I often tell students, “If you are confused, and think that nothing about this subject makes sense, it may be that you understand the topic.”  If you are confused, it may be that the subject is confusing rather than that you are stupid.  This is true of most of protozoan taxonomy today.  If you don’t understand the scheme, wait until next Wednesday and it will probably be different.

You are better to learn the 36 presently accepted phyla, get a general idea of the different classes of water creatures, learn the common species, then enjoy the pretty animals.  Often, just separating an acorn worm (Hemichordata) from an annelid (Annelida) or a thorny-headed worm (Acanthocephala), or distinguishing a Stentor from a Vorticella (and maybe a couple of species of each) is as far as most of us have time to go.

The reader is referred to David Goldstein’s Micscape article, “The Classification of Living Things” for further thoughts on this topic.  Those who want to feel much better about their humble understanding should peruse Walter Dioni’s well-researched article,  “An Annotated Key for Species of the Family Stentoridae and Two Related Families:  An Excursion Through the Taxonomic Maze” and muse on the torturous progression of the taxonomy of the simple Stentor.

If you are STILL feeling badly, carefully read Vd’acny’s recent (2013) paper “The Chaos Prevails: Molecular Phylogeny of the Haptoria (Ciliophora, Litostomatea).” and make sure that you take CAREFUL note of Figure 5.  Then consume a moderate quantity of hydroxylated ethane, go to bed, and pull the covers over your head for an hour.  When you emerge, you will feel better with the knowledge that much smarter people than you struggle with taxonomy and are busy messing up everything that you THINK you know, so why bother?


Dioni, W.  “An Annotated Key for Species of the Family Stentoridae and Two Related Families:  An Excursion Through the Taxonomic Maze.”

Ehrenberg, C.G.  Die Infusionsthierchen als vollkommene Organismen, Vols I & II.  Leipzig, 1838.

Gao, F. et al.  “The All-Data-Based Evolutionary Hypothesis of Ciliated Protists with a Revised Classification of the Phylum Ciliophora (Eukaryota, Alveolata)”. Scientific Reports No. 24874 (2016).

Howey, R. L.  “A Microscopic Loch Ness Monster.”

Jones, K. and Smith, M.  “Tear Of a Swan.”

Wikipedia.  “Christian Gottfried Ehrenberg.”

Wikipedia.  “Cytostome.”

Vd’acny, P. et al.  “Genealogical analyses of multiple loci of litostomatean ciliates (Protista, Ciliophora, Litostomatea).”  Molecular Phylogenetics and Evolution, 65, 397–411, Issue 2, November 2012.

Vd’acny, P. et al.  “Phylogeny and classification of the Litostomatea (Protista, Ciliophora), with emphasis on free-living taxa and the 18S rRNA gene.” Mol Phylogenet Evol. 59:510-22 (2011).

Vd’acny, P. et al.  “The Chaos Prevails: Molecular Phylogeny of the Haptoria (Ciliophora, Litostomatea).” Protist, Vol. 165, 93–111, January 2014.

Zhang, Q. et al. ” An Annotated Key for Species of the Family Stentoridae and Two Related Families:  An Excursion Through the Taxonomic Maze.”  Proceedings of the Royal Society B 29 Feb 2012.

Lacrymaria olor and the Litostomata – Beautiful Ciliates with Nasty Dispositions

How Bright is your Light Part IV: Thoughts on LED Safety

Since microscopists frequently spend hours looking at specimens, the question of the safety of LED illumination is significant. Concerns regarding the possible damaging effects of blue and ultraviolet light after conversion of scopes to LED light sources have been raised by several authors describing conversion techniques.  Traditional incandescent light bulbs produce most of their illumination in the red or far-red end of the spectrum, with modest output in the visible range and minimal ultraviolet production. Consequently, incandescent light that is comfortable to observe is unlikely to cause damage.  This is not necessarily the case with an LED light source, where shorter-wavelength blue light excites a phosphor to produce a second peak of mid to longer visible wavelengths.  The latter is less likely to be injurious to the retina unless the light intensity is high, but the UV component and the amount of primary-emission blue light are often poorly characterized.

The retina is a complex organ with ten layers, five of which form a complex, interconnected nerve network performing some basic image enhancement.  The arrangement of the retinal layers is exactly upside down from what logic would suggest:  the nerve cell layers are on top, and the photoreceptors are mounted upside down at the back of the retina.  Light must therefore traverse the nerve layers before activating the light-sensitive photoreceptors (images from Ted Montgomery’s site):

There are two types of photoreceptive cells in the retina: rods and cones.  Rods are the more sensitive and are responsible for low-light vision, but cannot distinguish colors, having only one light-sensitive pigment, or chromophore: rhodopsin.  This pigment, most sensitive to green light at 498nm, consists of a complexly-coiled protein that crosses and recrosses the cell membrane of the rod cell.  It forms a pocket for the pigment retinal, a Vitamin A derivative.  An opsin protein acts like a carefully-balanced tangle of springs;  when a photon strikes the retinal molecule, it undergoes a subtle structural shift (conformational change) that twitches the opsin into a slightly different shape. This  change also affects the  segment of the protein inside the cell membrane, where an active enzymatic site is exposed, resulting in a chemical cascade that causes the rod cell to depolarize and create a nerve impulse. This process is an important mechanism in cells where transmembrane signaling occurs:


Rhodopsin crosses the cell membrane seven times and contains the pigment Retinal  (red) in a small pocket.

Cones, the color-sensitive photoreceptor cells, are less sensitive, are the primary cells responding to daylight light levels, and have three chromophore pigments, L, M, and S, responding respectively at 564nm (Long wave, red), 534nm (Medium wave, green) and 420nm (Short wave, blue).  The normal human eye is therefore populated by L, M, and S cones.


The mechanisms of retinal damage by light are complex and incompletely understood, and the literature on this subject is both enormous and confusing (see Organisciak and Vaughan as well as Youssef et al).    Two broad categories of damage are recognized: thermal damage and photochemical damage.  Thermal damage reflects heat-induced retinal damage and can occur across a fairly broad range of visible wavelengths, but requires significant intensity.  At very high light intensities with focused spot retinal heating, shearing and mechanical effects from heating of the aqueous components can also become a factor.   Both effects are of concern with powerful sources such as infrared YAG lasers, but should not be a significant risk to the average microscopist.

Photochemical damage is of more concern; the risk for this form of retinal injury is concentrated in the shorter wavelengths (i.e., blue and violet) of the visible spectrum and peaks in the ultraviolet.  There also seems to be some photochemical damage that can occur in the midrange of the visible spectrum as well; this is likely to reflect energy absorption and subsequent chemical reactions by the chromophore pigments.

The human eye is protected from much of the effect of  high levels of visible light by the aversion response – pupillary constriction, head turning, and blinking.  In other words, bright light hurts, and we respond reasonably quickly, reacting protectively in about one-quarter second. This response offers considerable protection from light that we can see.  Unfortunately, the effectiveness of this reaction decreases significantly on both ends of the visible spectrum.  Infrared light passes readily through the corneal and vitreous; on exposure to a powerful Nd:YAG infrared laser, the patient may feel only a “pop” as focal explosive boiling creates a permanent blind spot.  Similarly, the aversion response to blue or violet light is weak, and the eye has no significant protective  response to ultraviolet wavelengths.

Research results and published opinions are contradictory.  Much work has been done in this area, as light-induced damage mimics many of the features of chronic degenerative processes such as macular degeneration, and chronic light exposure may be a factor in these processes.  Some studies have used very intense exposures and may not be good models for the effects of long-term, lower-level blue or UV light exposure levels.  In 2013, the US Department of Energy published a fact sheet on architectural light sources that stated “…LED products are no more hazardous than other lighting technologies that have the same CCT (color temperature)…”  Then in 2015, a French research team published a highly sophisticated study showing biochemical and structural damage in the retinas of rats exposed to overhead LED illumination.  Sources used included white and blue LEDs from Cree Industries and Nichia Corporation.

Ultraviolet light is almost entirely absorbed by the cornea, lens, and vitreous, producing lens cataracts, denatured protein swirls in the vitreous, and and the equivalent of a sunburn on the cornea. These injuries are clearly more related to high-energy sources like welding arcs than an afternoon spent watching the lazy movements of a hydra..  However, blue light can reach the retina, and it is the effect of the powerful blue peak pumping the phosphor that needs to be evaluated..  A recent paper from Taiwan showed changes in the mouse retina after exposure to “…Low-powered Family LED Lighting…

Cree, Inc. has done safety testing with its LED products and has concluded that its blue and royal blue LEDs can pose a hazard

To be continued…..


Note on learning new things:  I should really admit that, like much of my writing, I started off in complete and utter ignorance of all three of these topics.  Furthermore, I have never actually DONE an LED conversion of a classic microscope.  Like much of what I write, this series is a notebook of my learning process – a reflection of how I organize my thinking so that new information falls into order, like shiny ornaments on a well-designed Christmas tree. Since I’m doing all this work, I might as well write it down so others can share it.  I’m also driven by my own curiosity and sense of fun at learning new things that I can use on my own projects.  It should also be noted that complete ignorance of a topic has never stopped anyone from loudly expressing an opinion or writing voluminously about it.  If you doubt this, listen to any politician.


Building Technology Office, US Department of Energy, Solid State Lighting technology fact Sheet.  “Optical Safety of LEDs.”  Published June 2013.

Wikipedia.  “Rhodopsin.”

Zhou, X. E. et al.  “Structure and Activation of Rhodopsin.”  Acta Pharmacol Sin. 2012 Mar; 33(3): 291–299.

How Bright is your Light Part IV: Thoughts on LED Safety

How Bright is your Light Part III: LED Conversion

ledsfigure1 incr vibrance
LED Spectra (from Zeiss site)

In Part I of this series, we talked about a simple, home-grown fix for a defective, very basic illumination power supply on one of the most classic of classic microscopes, the Nikon S.  Then in Part II, we moved on to repairing or replacing the more sophisticated supplies  for incandescent lights on modern scopes from about 1975 onward, and met the ubiquitous switch mode power supply, or SMPS.  In this segment, we will review the ultimate, and perhaps best, solution for repairing or updating illumination on a classic microscope:  LED conversion.

Rather than rebuild an older incandescent light source, the most expedient solution may be to upgrade it to LED illumination that provides bright, white-light illumination with a durable system using little current, producing minimal heat, and using no expensive, hard-to-find replacement bulbs.  You can buy a readymade LED illuminator for some microscopes.  For many classic scopes, however, there are no off-the-shelf solutions, and you are faced with modifying the illumination system yourself.   This not necessarily a bad thing – in some cases, the conversion can be done fairly cheaply and easily using pre-built power supplies or a simple battery supply.


Many, if not most, modern microscopes have LED illumination systems.  Aftermarket LED illumination kits or adapters designed for a number of specific microscopes have recently become available.  A Google search for microscope LED illuminators and light sources brings up a variety of retrofit kits and illuminators, a number of which are generic, of Chinese origin, and may or may not be a good fit to specific models.  The most sophisticated yet cost-effective manufacturer of LED retrofit illuminators for older scopes seems to be retroDIODE LLC of Gardner, Kansas.  This manufacturer uses 3D printing to manufacture LED units specific for a number of the most popular classic scopes; this is their unit for the Nikon S-Kt:SKt 02SKt 01At $140.00 US on retroDIODE’s eBay store, this seems to be a reliable and reasonably-priced alternative to building an LED illuminator from scratch.  Promicra also makes some LED illumination systems, but no price is given, and the cost is probably significant.  ThorLabs manufactures very sophisticated microscope light sources in a considerable variety of well-characterized spectra for several major brands of scope, but the price is about $450-600 US, and they are only made for fairly modern microscopes.

Thorlabs Mid-Power_LED_for Zeiss
Thorlabs Mid-Power_LED_for Zeiss

Microscopes with simple, non-Kohler illumination systems are much less of a challenge, as there are a number of simple LED plate-type substage illuminators available on eBay for $20-65:

Simple Chinese LED Plate-style Illuminator - $65 on eBay
Simple Chinese LED Plate-style Illuminator – $65 on eBay

Similarily, eBay carries a multiplicity of ring LED illuminators for stereomicroscopes at very reasonable prices.

The main advantage of buying a commercial LED conversion kit is that you don’t have to mess with deciding on the best LED, choosing a power supply, using tools, tearing out the old system, or making the new system.  However, LED conversion kits are not yet available for all vintage microscopes.  Furthermore, with some ingenuity, a conversion can be done for very little money, and you then have the satisfaction of understanding your equipment better (and deciding what to do with all the money you saved).


Before converting to an LED system, one should really understand what a light emitting diode is and how it works:

The first emission of light from a crystal junction was recorded at Marconi Laboratories in 1909-7 by H.J. Round, using a gallium arsenide crystal with a “cat’s whisker” wire contact, much like a 1920s crystal radio.  The first true light emitting diode was created

(and its theory correctly described using Einstein’s new quantum theory) by the Russian scientist Oleg Lesov in 1927.   Lesov, largely unknown despite being one of the first semiconductor physicists, studied solid-state light emission and actually used primitive semiconductors successfully in amplifiers and radios.  However, in the age of

Oleg Losev, 1903-1942

vacuum tubes, the work of this unsung genius was largely ignored.  Caught in the siege of Leningrad in 1941, he tried unsuccessfully to get a paper on a three terminal semiconductor (possibly the first transistor) out of the besieged city, but the manuscript was lost, and he starved to death in 1942, along with a million other residents of the city.  Lesov spent much of his life working as a lowly technician, and his work was forgotten for twenty years.  Nikolay Zheludev and Tom Simonite have published fascinating notes on Lesov and the original development of the LED.

Siege of Leningrad

Most sources unfamiliar with Lesov’s work credit the invention of the LED to the independent efforts of four American research groups in 1962 (see Wikipedia article); subsequent intensive work by many groups led to the first commercially successful LEDs in the 1970s, and there has been an enormous volume of research and innovation since that time.

Simply explained, a light emitting diode consists of a junction between an N-type semiconductor material (with an excess of electrons in the conduction band) and a P-type semiconductor (with an excess of “holes”, or electron deficiencies, in the lower-energy valence levels).  Application of a current across the junction forces electrons and holes to cross the junctional energy gap.  The electrons and holes jump from higher to lower energy states, and the lost energy is released:

Energy levels across p-n junction in LED (from Wikipedia)
Energy levels across p-n junction in LED (from Wikipedia)

HOWEVER, not all such transitions, and not all semiconductor types, produce light.  Asking “…But…but…why?…” leads the reader into arcane theory and dark places that are better left alone.  Anyone who lacks a strong stomach and a knack for dealing with quantum entities that don’t really exist but still can go bump under the stairs, is advised to take two Dramamine, sign off the computer, and lie down in a quiet place.  The less wary will learn that the energy drop across the junction is referred to as the band gap;  gaps are classed as direct or indirect depending on whether the direction of the crystal momentum vector of the electrons and holes in the valence and conduction bands bands is the same or different, respectively.  If you don’t understand that, don’t feel badly – the crystal momentum vector is only a virtual vector, so it isn’t really there anyway.

Delving further, explanations of semiconductor junction physics descend into pages of little squiggly things that would look nice on a wall, but tend to produce headaches.  For those who would actually like to explore this topic further, the Wikipedia article on electronic band structure is quite good.

What this means in a practical sense is that some semiconductor junctions can produce light and some can’t.  Charge carriers (electrons and holes) in those with an indirect band gap go through an intermediate energy state (rather like a pinball bouncing around) and dribble their transitional energy away into the crystal lattice in the form of heat – this occurs with silicon and germanium.  With a direct band gap, charge carriers make only one transition, giving out a nice photon as each carrier makes the jump – gallium arsenide crystals do this, as do crystals of aluminium gallium indium phosphide.  The color of the emitted light can be changed by doping the semiconductors with impurities to adjust the width of the band gap.  The Wikipedia article on LEDs has a nice table of semiconductor materials and their respective spectra, as does the Zeiss site.

LED Structure (from ADLED site)
LED Structure (from ADLED site)

Once you can produce light from a semiconductor junction, how do you get it out of the junction and focused into a beam that is useful?  Actual LED chips are tiny, ranging from 1/10mm to 1.0mm in size.  Most commonly, they are mounted in a small reflective cup on one of two supports, the anvil, or cathode.  A fine metal contact wire from the post, or anode, connects to the surface contact of the LED.  These internal components are sealed in an epoxy case with an apical lens that concentrates the light reflected from the cup into a beam of predetermined width.

Unlike the simplistic pictures of junctions, the structure of an actual LED is quite complex, and reflects thousands of hours of corporate and academic research, published on thousands of pounds of paper, and wrangled over at hundreds of annual conferences.  Different manufacturers have different designs, and the average LED is a highly engineered multilayer device.

Modern LEDs may use Bragg reflector (dielectric mirror) layers, insulated layers that channel current flow around microscopic central etched cavities, surface grooving to decrease internal reflection,  and a myriad other microstructures, all aimed at increasing efficiency and decreasing light losses.  Terms like evanescent wave coupling are bandied about. All of these little marvels work in the Alice-in-Wonderland world of quantum devices, where the physical laws we are used to simply don’t apply.  A few samples give one an idea of the complexity and diversity of these tiny devices:

Grooves etched on LED surface decrease internal reflections and markedly increase light emission
Insulating layers channel current within the layers of the microchip
Different modes of light emission – lateral versus vertical from etched surface point source

These images provide only a taste of the many variations in LED design, and advances in LED micro-engineering are occurring on an almost daily basis.  Remember that all of this structural work is occurring in an object about the same size as the head of a small pin.

Note especially that the mechanism by which the LED produces light is completely different from that of the old-fashioned light bulb.  Incandescent bulbs emit light due to black body radiation; this kind of radiation results from the increasing vibration of atoms and molecules as bodies are heated to higher and higher temperatures.  Atoms have charges, and vibrating a blob of charged particles faster and faster as the filament heats up generates electromagnetic waves that we see as light, going from longer to shorter wavelengths as the atoms move faster.  The resulting light has a continuous spectrum that shifts from red to blue as the temperature rises:


Any body at a given temperature will emit the same spectrum; for this reason, the color of light bulbs or other light sources is often expressed as a color temperature.  Midday daylight is roughly equivalent to the light from a heated body at about 5600 degrees Kelvin.  Note also that black bodies like light bulbs are hellishly inefficient producers of visible light, with most of the energy output occurring in the invisible and useless, long-wavelength far reds and infrared.  Consequently, most of the energy from the batteries in an old-fashioned flashlight is wasted in producing heat.

For balance, the reader should also review David Walker’s superb article in defense of tungsten illumination.  His images of denser objects taken with near-infrared illumination are striking; these would be difficult to accomplish with the narrower spectrum of present-day LEDs.  His compilation of creative ideas for illumination on the Biolam microscope is also well worth reading.


Production of the daylight-balanced white light essential for most microscopy applications requires even more ingenuity, since LEDs typically produce only a single spectral band.  There are actually several ways in which LEDs can produced white-appearing light (summarized from the detailed but excellent Olympus Microscopy Resource Center article “Introduction to Light Emitting Diodes“).

Probably the most common technique is to combine a blue, violet, or UV-emitting microchip with a fluorescent phosphor.  The short wavelength, high-energy light from the microchip falls on a phosphor lining on the surface of the reflective cup.  The latter absorbs the short wavelength light, then re-emits the light energy (fluoresces) with a broad band of longer wavelengths:

"White" LED with phosphor-lined reflective cup
“White” LED with phosphor-lined reflective cup

Phosphors typically consist of an inorganic host matrix, often yttrium aluminum garnet (YAG), doped with a rare-earth element such as cerium.  The overall emission spectrum then consists of the primary short-wave (blue) band from the microchip, together with the longer-wave and broader emission band of the phosphor:


Other methods include LEDs where all of the emitted light comes from a mixture of phosphors and the blue source light is never projected from the device.  Such devices can use phosphors already developed for fluorescent lights; unfortunately, this method has a much lower efficiency than the mixed microchip/phosphor emission technique.

Instead of phosphors generating the longer-wave spectral elements, another group of LEDs in development employs a second semiconductor layer to absorb short-wave light and re-emit longer wavelengths.  This combination of a current-driven blue LED feeding an optically-driven, longer-wavelength LED is known as a photon recycling semiconductor, or PRS-LED.

In some cases, white light is generated by a three-LED RGB unit whose color can be varied by changing the current driving the red, green, and blue LEDs.  By balancing the intensity of each LED, any color can be produced.

HOWEVER, remember that, even though LED light may appear white to the human eye, it is still only an approximation of the continuous spectrum of white sunlight that floods Death Valley every afternoon.  White LED light is still cobbled together from individual, discrete bands, with peaks and valleys that are not present in the continuous daylight spectrum.  As such, supposedly white LED light may behave differently from true white light in certain microscopic techniques, or may render colors and stains differently.  Chromaticity considerations are discussed in the Olympus LED reference.  This site also has a good general discussion of visible light sources.  The Zeiss web site also has a thorough discussion of light emitting diodes in microscopy.

However, when these theoretical concerns are put to practical testing, as in David Walker’s excellent side-by-side comparison of slides viewed by tungsten light and illumination from a Phillips Luxeon III LED, the images looked almost identical, with the LED colors actually being slightly more vivid.  Mr. Walker also discusses the significance of the color rendering index (CRI) in older and current LEDs.  To my pathologist’s eye, the color differences are slight, and seem functionally insignificant.

Non-Koehler illumination systems are reasonably tolerant with respect to the light distribution from an LED source.  With Koehler illumination, the angle at which the cone of light from the LED spreads, the evenness of the light distribution, and the presence of shadows from small internal structures such as contact wires all become critical.  As part of its thoughtful discussion of microscope illumination, John Walsh’s Micrographia site has a discussion of features to look for in an LED with respect to microchip size, as well as the curvature and positioning of the lens:


Unless you break it, an electric light is really a pretty simple-minded and amiable creature.  The more electricity you feed it, the brighter it gets.  You make it brighter by increasing the voltage applied across the filament, and if you plot the current flowing through (and heating) the filament, it follows the applied voltage in a regular and predictable manner:


(From I B Physics Stuff)

The graph would be a nice straight line if it weren’t for the fact that the resistance of the filament increases as it heats up.

John Powell aptly describes what happens if you try this experiment with an LED:

“…The relationship between current and voltage in an LED is non-linear. As the voltage increases from zero there is only a trickle of current and no noticeable light. At about a volt and a half … the current begins to increase appreciably and the first glimmers appear.

LED current graph

At two volts the LED is bright and with a fraction more it’s very bright. Once over about 2.2 volts, the current rapidly soars beyond safe operation. The LED soon overheats and dies…”

The reason for this behavior is that the applied voltage increases until it forces electrons and holes over the energy barrier at the junction between the two different kinds of semiconductor (the n and p layers).  Initially, nothing much happens until this threshold energy is reached.  However,  once the cascade of electrons begins, small increases in voltage result in a deluge of charge carriers across the junction and, like water eroding a dam, the LED can quickly be destroyed.  Consequently, LEDs cannot be dimmed by increasing the voltage; instead, the voltage must be held relatively constant and, unlike incandescent bulbs, dimming is accomplished by varying the current through the LED.

The water-over-the-dam analogy is really quite good.  With the LED, imagine you’re filling up a dam; the water level rises and rises, yet nothing much happens.  When the water reaches the top (junctional energy barrier), a trickle appears over the edge.  A bit more, and there’s a nice flow down the spillway.  A tiny bit more, and we have a torrent over the top that eats away at the dam and quickly washes it away.  To control the amount of work done by the water behind the dam, we use the gate on the spillway to control the flow (current).  Within limits, the water level behind the dam (voltage) can rise or fall, and we just open the spillway more or less to compensate.

An electric light behaves more like water running through a V-shaped canyon – raise the level or pressure (voltage) at the beginning of the canyon, and more and more water flows through.  The water level rises in the canyon as flow increases, but it is a reasonably steady and linear process.  You will finally reach a point where the waters wash away the walls of the canyon or overflow across the countyside (the point where the lamp burns out,) but the flow has been a fairly regular progression up to that point.

There are two ways of supplying power to an electronic gizmo: through a voltage source or through a current source.  Most sources of electric power with which we are familiar are voltage sources:  our 110V or 220V AC wall plugs, a 1.5V flashlight battery, etc.  A 12V car battery can run a flashlight bulb for a week, or the starter on your car for a minute.  In both cases, the voltage stays fairly constant at about 12 volts, but the current varies from a tiny trickle with the bulb to a hefty 100 amps for the starter.  Putting the wrong gadget across the battery results in a shower of sparks (and a melted gadget) as the current soars and burns it out.  This is a good example of a voltage source – the voltage stays pretty constant, but the current can vary enormously.  This is how we usually think of electricity – we decide the voltage, and the current is just whatever is needed, up to the maximum capacity of the source.  BUT – this is NOT what we need to keep our LEDs happy and unfried.

We are not used to stabilizing the current first and letting the voltage follow along;  this is, however, the nature of a current source – a power source that provides a constant current regardless of the voltage in the circuit.  Note that many solid-state power supplies are pretty happy with a range of voltages, and you can plug  many electronic gadgets into 110 or 220V and they don’t seem to notice the difference.   An electric welding power supply is a good example of a constant-current source, where the amount of metal deposited in the weld is a function of the current.  Similarly, LEDs need to be powered by a source that, at the minimum, keeps current flow from destroying the diode, and ideally, allows the current to be varied in a stable and regulated fashion in order to control the light intensity.

The simplest and most primitive way of creating a current-controlled power source is to take a voltage source (such as a battery or standard power supply), and add a series resistor in the circuit.  This does not create anything like a well-controlled source of amperage, but it does limit the maximum current that can flow through the LED, although in a way that wastes power.  As John Powell notes:

“…The usual approach is to put a resistor in series with the LED. The combination is still non-linear, but in a much more well behaved manner. In fact, over the range of safe operating current, it acts incrementally linear.

LED with resistor

The catch is that it wastes power. If a 12 volt supply is powering a single LED-resistor combination, 2 volts goes to the LED and 10 to the resistor. Only one sixth of the power makes it to the LED.”

However, with an LED, the total amount of power used is small, and resistor losses may not be important.  The following is a simple LED current limiting and dimming circuit  involving only a resistor and potentiometer, working from a 5V power supply:


(From Phil Frost, EE Stack Exchange)

The disadvantage of these very basic circuits is that they are inefficient and may not provide uniform dimming as the potentiometer is rotated.  Better current control, uniform dimming capability, and less power wastage require a more sophisticated circuit.  Searching the literature, as I did for a week, reveals a mind-numbing collection of complex circuits using operational amplifiers, integrated circuits with optical transistors, Zener diodes, etc., etc.  Fortunately, when one understands its relatively simple needs, the LED is actually a fairly amiable and undemanding critter that is happy with a ham sandwich for dinner every evening, and doesn’t require champagne.  However, it is a major project to find a simple, constant-current supply circuit with dimming capability for a single LED or small group of LEDs; many of the available circuits are meant for constant intensity lighting circuits or are designed for residential lighting.

Frank Weithöner (see below) describes a simple variable current two-transistor circuit for powering and dimming a microscope single-LED illuminator:


The two transistors, the MOSFET BD237/243 and the general purpose NPN BC546, are at present commercially available, and conversion datasheets are readily available for equivalent transistors.


Let’s now turn (literally) to the nut and bolts of doing your own LED conversion.  If you decide to design and build your own unit, there are a few references available on the internet.  However, there are as yet no standard techniques for retrofitting incandescent illumination systems with LEDs. LED illumination systems are relatively new and rapidly evolving for all applications, including automotive and residential, and articles on retrofitting classic microscopes are scanty as of 2016.

One important and as yet unsettled question is the number of LEDs in the light source.  Where will the microscope industry land – with a single, high-intensity LED and lens, where one deals with the same problems of uneven center-to-edge brightness found with conventional lamp filaments, or with a multiple-LED source, which is more even between center and edge, but where one must strongly diffuse and evenly spread the light from several discrete sources?  So far, the replacement illumination systems for binocular scopes with or without Kohler illumination capability have used high quality, high intensity single LEDs with focusing lenses.

One new development that may have promise for producing a very uniform light source without the problems of evenly spreading light from a point source LED is COB, or “Chip-on-Board” technology.  In this new design, multiple small microchip LEDs are arranged in an array to form a small, uniformly illuminated lighting panel.  While many of the manufacturers are in China, American lighting corporations such as Cree Inc., headquartered in South Carolina, list a wide variety of small (1-4 cm) lighting panels using this technology.  This new light source is only recently commercially available and has not yet been used in microscopy but has, I think, considerable promise.

Cree CXA2 1.2cm COB LED

As of June, 2016, there are no online references on the possible uses of COB technology in microscopy.  One question, to which I do not immediately have the answer, is how a small flat-panel source fits in with Kohler-type illumination and maximizing resolution.

There a few articles available on LED conversion of older or semimodern microscopes, all of which to date have employed single-chip LEDs with focusing lenses. One excellent site is Frank’s Hospital Workshop, devoted to hospital equipment maintenance in impoverished Third World countries. Frank Weithöner describes conversion of Olympus CH-2 microscopes to LED illumination using a single Luxeon Star/O LED with lens.  He also has a good general page on power supply basics.  This is one of the most carefully crafted sites that I have encountered, with well-written, practical advice on the maintenance of a variety of medical equipment.  The images below, from Mr. Weithöner’s site, show an Olympus CH-2 microscope before and after LED conversion.  The small SMPS 5V power supply is on the left in the converted base, while the potentiometer and current control circuit are at the top:



If the standard power supply can be modified, or a tiny converter supply built into the socket, this might allow conversion to LED without significantly altering the microscope.  A similar mounting of an LED onto a microscope bulb base has been described by David Walker.

The above sites provide details of professional-level conversions.  There is also a place for simple and inexpensive solutions using local materials, similar to spending $5.00 on a cheap dimmer to rebuild a Nikon S illumination system (see Part I).  The cheap and readily-available LED flashlight may prove a simple fix for illumination problems.  When my Nikon S illumination system failed, I removed the lamp and inserted a cheap (10 for $12.00 at the thrift shop) seven-LED flashlight that fit into the lamp housing.  Without a good diffuser, it was nothing that I would use for photography, but it worked and the scope was usable on an emergency basis.

A very simple conversion of an American Optical 150 microscope into a portable scope, using an off-the-shelf LED powered by a 9V battery, was published in the November 2011 issue of Micscape magazine by Bill Resch.  The LED was clamped in place and supported on its two wires in the same place as the lamp filament. Bill’s published images suggest that the illumination was quite even.

In the March 2011 issue of Microbe Hunter magazine, Suphot Punnachaya described a simple LED conversion of a Chinese microscope’s halogen illumination system using an LED array obtained by sawing off the front inch of an inexpensive LED flashlight.  Power was supplied by an old cell phone 5V “wall wart” charger connected to the LED through a series resistor and rheostat.

An even simpler conversion of a Zeiss Gfl scope was describe in the August 2013 issue of Micscape by Franz Schulze; this used a $2.00 single-LED flashlight with lens, purchased from the Dollar Store.  The flashlight was mounted in a simple wooden sleeve machined from a wooden dowel, and placed into the port for the lamp assembly.

It should be noted that microscopes such as the Zeiss Gfl and the Nikon S, where the bulb assembly is inserted  into a port or sleeve in the body, are easiest to convert to LED illumination.  Those models where the bulb is enclosed inside the base require somewhat more work, ingenuity and removal and remounting of  components.  In these cases, assembly and mounting within the base might be further simplified by using a hot glue gun to mount components (or the LEDs themselves) rather than machining clamps.


Powering the LED illuminator can be as simple as using coin, flashlight, 9 volt, or rechargeable batteries, since the power requirements of many LED sources are low enough that batteries are a viable option.  The need for a series resistor is described above.

To power an  LED light source from 110 or 220 volt mains AC, one has the same options as for the incandescent power supplies discussed in Part II:  buy a small LED power supply (known as an LED driver) or build your own circuit. Many inexpensive dimmable LED drivers are available on the internet; they can be wired with a simple Triac dimmer to control the input voltage and hence the output current:


Small Triac dimmer – disconnect the potentiometer connections and reconnect it with three wires, then use it to replace the intensity control


Small dimmable LED driver supply – under $20 online from many sources

Specifications for this dimmable LED driver

 This is probably the simplest option for setting up an LED supply within the base of a microscope.  One might consider combining the small dimmable LED driver with the guts of a cheap dimmer, the latter replacing the microscope’s illumination control and the rest of the dimmer’s circuit components being transplanted as a unit, as described in Part I.  However, you might be better to buy a small dimmer unit matched to the LED driver from the same supplier.

As far as building your own LED power supply from scratch, you can do it if you are more familiar with modern electronics than I am.  Yet in these days of mass-produced circuit boards, this seems like unnecessary complexity. Browsing LED driver circuits online resulted in many pages of complex diagrams using integrated circuits, MOSFETS, small inductors, etc., etc.  I have a PhD, and I felt lost and frustrated.  This seems like a lot of work when I can spend $10-20 and buy a smaller and better-made driver from any one of fifty electronic supply houses, then add on a simple dimmer circuit and figure out how to situate it all within the particular configuration of the microscope base..

One  more thought – the power supply for your microscope’s new LED system may be sitting right in your desk drawer.  Or in a box your neighbor’s garage.  Or at your local thrift store.

PS 02
Wall of “wall warts” at Goodwill store

For the last twenty years, orphan power supplies for all kinds of outmoded electronic gadgets have been piling up in boxes and drawers and finding their way to thrift shops.  Your local computer service store may have a box of disused power supplies in their back room in various voltages.  Some of these are wall warts, while others are inline supplies.  These originally powered computers, printers, cell phones, zip drives, makeup mirrors, cordless phones, games, and multiple other devices that need low voltage DC from a wall socket.

Simple, old-fashioned “linear” transformer-based wall wart

Now, remember that these small power supplies are voltage sources and are of two types: SMPS and linear.  Most of the newer supplies will be switch mode supplies (SMPS) that convert, reconvert and regulate their low-voltage output, with light, high-frequency transformers (see SMPS discussion in Part II).   The older, bulkier units are usually very simple 60Hz linear supplies, with bulky, low-frequency 60Hz transformers.  You can tell the difference by just hefting them in your hand; SMPS are very light, while transformer-based linear supplies are quite heavy.

To convert either to a usable current-based power sources, one can employ either a basic resistive network or a simple current regulating circuit such as that described by Frank Weithöner (see above).  It may also be helpful to Google something like “LED supply from wall wart.”  This will bring up multiple posts with ideas of varying usefulness from electronic hobbyists.  John Bryant’s “A Dummies’ Guide to Working With Wall Warts” is helpful, though it focuses more on voltage than current sources.  I have not cited any others specifically, as they change on an almost daily basis, but they can have interesting ideas.

Be aware, however, that some of the cheap, no-name wall warts lack the protections and sophistication of brand name devices and can be dangerous (see Ken Shirriff’s article “Tiny, Cheap, and Dangerous“):

Genuine iPad wall wart versus cheap copy.  The fewer components in the fake iPad supply reflect the lack of regulating and protective circuits.  Notice that the large ground post in the copy is probably plastic and, unlike its metal counterpart in the real iPad supply, does not seem to be connected to anything.

Do not dismiss the big, heavy, 1980s-1990s wall wart.  Though they are older, heavier, and clunkier, the old linear wall wart of any brand with its heavy transformer is probably safer than any cheap modern SMPS supply.  The big transformer serves as an isolation transformer, preventing the user from directly connecting with line current and risking a serious shock.  A good SMPS is a complex device with safety and isolation features built in, but a cheap switch mode supply with all the safety circuits left out, no ground, and possible internal shorts will still function, and you can’t tell the difference from outside the case.

Whether you use a purchased, ready-made LED driver or put together a supply from inspired scrounging of bits and pieces at the thrift store, this can be an inexpensive but creative project.  LED illumination technology is in its infancy, and new products are becoming available every day.  Consider the conversion a challenge to your inventive talents! Or, if you just want to plug in a storebought box with a cord on one end and an LED on the other and get back to chasing Peranema, that’s fine too.  If either approach helps you to fix your dead scope or restore a classic beauty, it’s worth it.


Note on best general references:  The Micrographia site has a good general discussion of microscope illumination, including thoughts on optimal LED design for microscopic illumination.  The ZL2PD Amateur Radio web site has an excellent basic discussion of switch mode power supplies.  The Olympus Microscopy Resource Site is a very good overall resource on microscopy, with a superb article on all aspects of LED function.  The Zeiss site Fundamentals of Light-Emitting Diodes (LEDs) is also very well-written, informative, and dense with information, especially with respect to LEDs’ potential role in fluorescence microscopy.  I have tried here to summarize just the information necessary for a general microscopist’s working understanding of this new technology.

Part IV deals with the physiology of the eye as it pertains to the many unresolved questions regarding LED safety.


Davidson, M. W.  “Fundamentals of Light-Emitting Diodes (LEDs).”

Elliot, R.  “Electronic Transformers, the Good and the Ugly.”  Elliot Sound Products website, accessed April 4, 2016.

Engdahl, T.  “Light Dimmer Circuits.”

Frost, P. Posted on Electrical Engineering Stack Exchange.  “Using a Variable Resistor to Dim an LED. “

IB Physics Stuff.  “Electric Circuits.”

Intelligent Controls pamphlet, accessed March 25, 2016.  “How a Dimmer Works.”

Kuphaldt, T.  “Fundamentals of Electrical Engineering and Electronics:  The Triac.”

Majumdar, S.  “How Switch Mode Power Supplies (SMPS) Work.”

Micrographia Site.  “The Microscope Lamp.”

Powell, J.M.  “LEDs and Dimming.”

Spring, K. R., Fellers, T. J., and Davidson, M. W. , Olympus Microscopy Resource Center. “Introduction to light Emitting Diodes.”

Weithöner, F. “Microscope Conversion to LED Light.”

ZL2PD Amateur Radio Website.  “Introduction to Small Switchmode Power Supplies.”

How Bright is your Light Part III: LED Conversion

Seattle: The Tiny Desk Aquarium

On one of my many trips to Seattle, I began contract work with the University of Washington, and decided to start a new aquarium in my hotel room.  For $2.99, found an 8″ cubical vase with thin, fairly even side, and stocked it with plants and mud from the U of W lakeside grounds and in ditches around Koll Business Park.


Anyone who has been around marshes knows that oily films on the surface are very common, and result not from pollution, but from substances released from decaying organic matter. Examining these quickly shows that they are a rich and possibly poorly explored environment.

The tiny hotel desk aquarium is doing well, but has developed a film of this type as the leaves, organic debris, and mud stirred up during collection undergo natural processes of degradation; the film can be best appreciated by comparing it to the clear avenue in this image (see image where you can see a clear avenue in the film):

Aquarium with Surface Oily Slick
Aquarium with Surface Oily Slick

Passing a slide through this film brought up a layer of mucky brown substance:SURFACE SLICK ON SLIDEYet the microscope revealed an area of almost unbelievable richness of bacteria, flora and fauna, as shown in the two videos, one of the film of degenerating plant material, and one of the water between islands of decaying fibers. Resolution is not perfect, but best I can manage in the field, and you are looking through a fair anount of guck on many of the levels. I count long strands of acinetobacter, gliding algae, cyanobacteria, ciliates, sessile algae, a rotifer, a small water flea (not shown), Synura or a similar species, several Vorticella, and many flagellates, a few of which I think are euglenoids:

Many questions come up regarding the function of these surface films and their role in the marsh in terms of gas exchange, absorption of sunlight, breakdown and recycling of organic material, possibly aided by solar energy, etc, etc. Fascinating – like having a webcam into a stretch of Amazon rain forest!

Seattle: The Tiny Desk Aquarium