Laboratory Dodges and Wheezes
(not to mention war stories)
Resin or balsam film for arranging slide subjects
Glass beams for pointers and balances.
Dead man’s switches & other precautions.
Introduction
It is many years since I last entered a laboratory in anger, so to speak, but I remember those days with pleasure. Laboratory dodges and the development of competence (and, more hazardously, confidence) were the sort of thing that tempted one into science in the first place. In those years everyday laboratory discipline was still developing as a professional concept, and safety was still considered a matter of technique and common sense, rather than equipment and book rules. You didn’t generally wear a mask or gloves unless there was some special reason. Some didn’t even wear a white lab coat, on the principle that if you knew your stuff, no lab coat was necessary.
I don’t say that the modern rules are bad, or that you shouldn’t have safety as a primary concern, but I do often see youngsters who simply do not understand what some of us called laboratory etiquette: how to carry a burette, and why not hold it in the convenient horizontal position, even when it is empty and dry, how to handle a stopcock and push a glass tube through a cork stopper etc. Those are skills, and their loss, though unavoidable for most laboratories, still is a loss.
But that is not the reason I wrote this essay. I just had developed (as we all do, else what on Earth are we doing in a lab or in the field) some of my own tricks, and if they are not all of value to many readers, at least some might be entertaining. They come without much recommendation, and with no warranty of either safety or results, but I hope you enjoy some of them.
Rubber tubing still occurs in most labs, though much equipment now comes with everything prefabricated and assembled. However, there used to be more tubing, and more varied in bore, wall thickness, and type of rubber. Typically it was white latex rubber, or red, or black. The type I speak of here, did not have woven fibre reinforcement in its wall. One of the routine nuisances was the difficulty of neatly cutting tubing of more than a few millimetres to the required length when required. One could, if suitable shears or guillotine were available, chop through the tubing more or less neatly, but the result was not impressively smooth, and such tools commonly were not at hand. The usual practice was to saw through the rubber with as sharp a knife of blade as might be available. Lubricating the work with soap or glycerol (not oil!) improved the ease of the process and the quality of the work, but it was messy, and only a skilled operator could do so without irregular knife marks, not only ugly, but often unsuited to one’s needs, especially if the tubing was very thick.
Messing around as I often did, I discovered this algorithm:
- Mark the position where the cut was desired.
- On either side of the mark, grip as short a length of the tube as might be practical; if there is no helper available, grip at least one side with a vice.
- Stretch as the tubing as tightly as practical; the mark will stretch too. Sever the tube at the middle of the mark.
- Using as sharp a blade as might be available (A well-maintained knife, or a handyman’s knife with disposable blades will do) cut the stretched tube with a single strong, slicing, chopping motion. Nothing fancy, no sawing, just a single slicing blow. If you have stretched it tightly enough, the tube parts in one snap, almost like snapping a scored glass tube. A successful cut leaves a mirror-like surface, so smooth that it is practical when one wishes to mate the cut surfaces end to end, to hold them together in any way convenient. They will fit without leakage if the internal pressure is not excessive. The quality of the cut puts a guillotine to shame. The technique works with some kinds of stretchy plastics as well, but not generally with reinforced tubing.
Lubricating rubber and glass
For lubricating rubber and glass, such as when trying to force glass rods or tubes into rubber tubes or stoppers, soap solution or glycerol or glycerol jelly, are easier and safer than trying to work dry, and less messy and harmful than oil or grease. For thin latex, such as condoms, oils are downright hazards, and cause rapid failure, so they should be absolutely avoided.
Forcing rubber tubes onto rods and rigid tubes.
A perennial chore in labs that do not buy all their kit ready made, is the fitting of rubber tubing onto the end of rods and tubes; if the tube is big enough to go on easily, it fits too loosely to be satisfactory; if it is tight, it stubbornly resists pushing.
The usual approach is to lubricate the tube with something like soap or glycerol. That helps a lot, though not enough to banish the problem except in marginal cases. A useful dodge is to connect the other end of the tube to a source of compressed gas or water of sufficient pressure (suitable mostly for outdoors, as there is much splashing and squirting, unless you work under water or inside a cabinet or shower: then block the far end of the glass tube if that is the object (a rod is already blocked)). Connect the rubber tube to the source of the gas or water and push the glass into the tube. The pressure forces the end of the rubber tube open, balloon-style as soon as it is blocked. Lubrication may help, but is not always necessary. When it works, this is a quick solution.
When it is practical, it is less effort to dip the end of the rubber tube into a suitable volatile solvent for a few hours, perhaps overnight, depending on the nature of the material, and on how thick the tube wall may be. I used to use benzene or dichloromethane, though neither is popular nowadays, both being regarded as health risks. Toluene might be safer, less toxic, though it still is a fire hazard. In a few hours the tube swells, and you can get quite a thick item into its opening without difficulty. You even can use the trick in joining a rubber tube to a thicker tube that way.
Leave the joint in a warm spot for several hours for the solvent to evaporate. If lab organisms are to be exposed to the solvents evaporated from the swollen tubing, do not use it before ventilating it in a warm spot for several days
The swollen tube is quite fragile until it has shrunk to its original size, so if you need to use force on it, be gentle.
Or think again.
Viscometry
At one time I was doing some research on aerial spraying, and needed to measure the viscosity of the fluids to be sprayed. We were not physicists, and we had no viscometers, not even the primitive Ostwald viscometer: a glass U-tube with a bulges at intervals to contain the fluid being investigated. You sucked the fluid into the reservoir bulge, and measured how long it took the fluid to flow down again through a capillary tube. Then you calculated the viscosity according to the flow times.
I had no such kit, no experience of viscometry, just the lab resources of a government biological lab. And a pretty good library. Available equipment included constant temperature rooms, glass and rubber tubing, tap-driven water-jet suction pumps, glassware, and a good range of fluid lab chemicals.
It was a case of create, or do without.
Start with a good, straight length of commercial so-called capillary glass tubing of perhaps 10 mm diameter with an internal diameter of perhaps 2 mm. Choose or cut a convenient length, possibly a metre or so, from a good, straight specimen of such tubing. To use it in creating real capillary tubes, one heated this raw tubing and drew it out to the desired thinness. That worked very well; it could produce almost microscopic tubing of good quality; however, for our required viscometry the raw tubing was just about right. Of course, for microscopic purposes, or measurement of viscosity of minute volumes of fluid, there was no reason not to draw it into much smaller diameters, but we had no such need.
First bend the tube into a right angle some 15 cm from the end. The short, straight arm becomes the viscometer’s connection to the fluid in the open beaker acting as the test fluid reservoir. Score a mark round the tube to show how deeply it should dip vertically into the reservoir beaker holding the sample fluid.
Next, at equal intervals of about 15 or 30 cm, score visible markings round the long horizontal shaft, so that one could see when the meniscus of a test liquid passed relevant spots. I made do with two intervals, though three should in principle have been better.
Then bend up the free end of the tube to form the suction end so that it points up at a convenient angle, say about thirty degrees. Terminate it with a short length of utility red rubber tubing. The length of the suction end hardly matters; its only function is to connect the viscometer to the suction device to draw up the fluid from the reservoir beaker. Into the tube. The rubber tube had been cut cleanly as described in the previous section, so that by lightly pressing the two ends of tubing together they formed a sensitive and convenient, not to say foolproof, means of sucking up the fluid into the working tube.
Fasten the viscometer horizontally into a couple of laboratory stands. The sample input end points vertically down into the reservoir beaker, with the suction end conveniently near to the pump. Use a level to make sure that the horizontal part of the tube is truly horizontal. Suction can be applied by pressing the vacuum tube mouth against the tubing on the viscometer. This may be required either for charging the apparatus for a reading, or for sucking in a rinsing fluid or drying air as required.
When ready, and with suitable commonsense precautions concerning temperature and contamination etc, start the suction and apply the vacuum tube to the rubber tube end. Suck till the fluid passes up and horizontally past all the scored markings. Remove the suction source and be ready with the stopwatch to record how long the fluid took to pass the scored makings on the horizontal section. I took measurements of various standard fluids at various temperatures; the times required ranged from less than ten seconds for the likes of ether, to several hours for some cold oils of interest.
Record the times at which the meniscus of the outflowing fluid passes the various scored markings on the way to the reservoir. Standardise the viscometer by first doing timing runs with fluids of known viscosity at suitable temperatures; there are tables in the sacred Handbook of Chemistry and Physics, and no doubt online too, but in those days there was no such thing as online; access to even a mechanical desk calculator was a rare major luxury.
Calculate the various results accordingly, and graph the viscosity readings of the fluids. On suitably logarithmic paper (no computers, remember!) the lines I drew were so straight for all the fluids that I tested, that one could hardly believe them. The speed of measurements was an order of magnitude faster than items of Ostwald equipment that we borrowed briefly from a neighbouring institute to confirm that our results were reasonable, and our single device served for a far wider range of fluids than the Ostwald did. It also was cheaper, and easier to use and to clean and required far less sample fluid. I could have drawn finer tubing, but was already using such small volumes that there was no need to do that.
Improving on killing bottles
In entomological field work it so commonly used to be necessary to catch insect specimens for collections that “killing bottle” was a standard term. Killing insects for collections is no longer freely permitted in most countries or universities, but something of the type still is necessary on occasion. The classical killing bottle had a source of poison, usually in the bottom of the bottle, soaked into a suitable spongy item. It tended to be messy, or dangerous depending on the substance, and if it did not rapidly calm or kill the insect, captured specimens tended to damage each other or themselves. The problem was particularly bad with moths and butterflies, and even if the specimens were not visibly damaged, some killing agents tended to cause them to stiffen as they died, making it very difficult to set them up as usable specimens.
Since those days we have found that carbon dioxide, though it does not kill most insects, does rapidly send them to sleep. Taken back to the lab within a few hours, they can be released if it turns out that they are not needed as specimens, or they can be killed painlessly by freezing if specimens are required. How best to keep CO2 in a killing bottle in the field, sometimes for days, is not always easy to tell, but it can be worth it. A particularly convenient means is to store a lump of dry ice in a vacuum flask or similar vessel; this both anaesthetises and freezes the insect very rapidly.
What is more, when such an insect is thawed after many hours of freezing, it generally is relaxed and easy to mount properly for preservation; in fact, as long as it is kept frozen before mounting or preservation in suitable fluid such as ethanol, its DNA too, is well preserved instead of being largely degraded at ambient temperatures — in fact, if the DNA is required, such a specimen, if still frozen, can be placed under vacuum and freeze-dried. Many other killing agents leave the insect stiff and hard to work with, often requiring special techniques that spoil the specimen.
Freezing also is a cheap and humane means of killing insects in the lab, and not as messy as squashing or poisoning them. I still shudder on having read of the way in which naïve biologists of a few decades or centuries ago killed specimens by dumping them in alcohol: one case was when someone caught a large mother spider covered with babies and wished to preserve them. He separated them and dumped the mother into alcohol; she was still struggling some time later, but the assumption was that she was dead and the struggles were reflex movements. So he dumped the babies in after her.
She recognised the babies and gathered them to herself in a ball and remained like that till they all were dead, whenever that might have been. I understand that he was badly shaken, as indeed I still am today when I think of it. Intelligent freezing would have avoided the incident, though I must say that I would have needed the specimen badly before I killed such a mother with young. (OK, OK, so I’m squeamish — make the most of it.)
Pseudo Air Leaks
For many purposes one needs to boil a liquid off a solution, and often high temperatures are not acceptable, because they would destroy or degrade the substance being concentrated. In such cases one often does the boiling at reduced pressure, which lowers the boiling temperature. However, that often leads to severe “bumping”, sudden blasts of violent boiling that cause spillage or contamination or damage of equipment or loss of the material. One would expect the addition of boiling stones to prevent that, but some solutions simply bump worse when one adds boiling stones, and some boiling stones lose their effect once they are saturated with the liquid.
A powerful technique is to insert a capillary tube through the stopper and down to near the bottom of the boiling vessel. This is known as an “air leak” and it is cheap and effective when appropriate, because the tiny stream of bubbles infallibly nucleates bubbles to keep the boiling liquid from superheating.
However, for some liquids one cannot accept bubbling air through, because the oxygen would harm the material in question. For this I find a pseudo air leak to be useful. One could simply attach a source of nitrogen or argon to the capillary input tube, but that is a finicky thing to do without breaking the fine tube. Instead I prefer to select a reservoir of a harmless fluid that has a much lower boiling point than the liquid to be boiled off. Call this the "leak fluid". For most purposes ethyl ether is a good leak fluid, but many others are more satisfactory for some applications. Examples include pentane, methylbutane, chloroethane and so on. (If you don’t know which will suit your needs, then you should not be doing any of this anyway.)
The trick here is to prepare a capillary tube of suitable dimensions, draw its point to as fine a hole as is practical, and blow a bulb of suitable size into the other end. This bulb will act as a reservoir for the fluid to pass through the boiling liquid. Curve the top of the tube so that the reservoir hangs erect. It is not in principle necessary to draw the long shaft of the tube to capillary dimension, only the point, so the difficulty of inserting the tube through the stopper can partly be avoided. Prepare the vessel stopper so that you can pass the capillary through it, and seal the hole to fix the capillary in place.
To charge the reservoir with the leak fluid, warm the reservoir while holding the point in the leak fluid, then let it cool till it has sucked up some fluid. Heat this in turn and cool it again. It might take two or three heating and cooling cycles to fill the reservoir as desired. Depending on the fluid, it might be necessary to charge the reservoir in a refrigerated room. Cool it finally, and set up the apparatus for the boiling.
It works smoothly.
One often needs a roughened or fogged surface on glass or glazed tiles, vessels, or microscope slides, whether for friction, light diffusion, writing on the surface with a pencil, marking, or more other purposes than I can think of offhand. This trick also works for etching hard metals in such a way. In particular it can be useful when working with very fragile objects; decades ago there used to be an ad in the Scientific American, showing a light bulb sliced like an onion by a commercial device. But if you have a source of air pressure and a milling device (such as a ball mill) and a source of sharp sand, broken glass, or fine abrasive powder, then a home made miniature grit-blaster may be particularly helpful.
If the pressures permit, it is best to make the grit vessel of glass so that you can see what is happening inside, but for heaven’s sake make sure that everything that is subjected to pressure is in a mesh cage to prevent some very ugly injuries when it bursts, as it certainly will, sooner or later. Also wear both eye protection and a mask to avoid breathing fine particles; fine mineral powder is dangerous, treacherous, and incredibly insidious. Don’t come crying to me if you neglect these warnings! This is one of the cases where experience is a poor teacher: it presents the exam before presenting the class.
OK. Find some place to work, preferably outside; such powder is not only dangerous, but messy.
The principle is that if you put fine, loose, dry abrasive grit into a suitable vessel, and blow it at out at high speed against a smooth, brittle surface, such as glass, it will etch the surface. Various versions of the idea have been used for various purposes since at least the late nineteenth century, but one seldom sees it used in labs, especially not home-made.
Put the dry grit loosely into a suitable vessel with a stopper that will withstand the pressure, leaving lots of free space above it, with an input tube that does not quite reach the surface of the pile of grit, then the incoming stream of air will create a miniature sandstorm. The output tube should end much further above the pile of grit, so that it only gets finely separated grains without risk of clogging. It should lead into a flexible output tube that you can direct at the target surface. The bore of the output tube should be at least ten times the size of the greatest grit particle.
Finely ground glass or grit can be blown against a smooth surface, to produce almost invisible pitting, but you also can mask the surface with masking tape or by writing or drawing on the smooth surface with a solution of rubbery plastic or wax. That is enough to prevent pitting, and you can mark the pitted surface with pencil or something similar. You also can melt glass or glaze and blast the molten surface with coloured glass or salts that melt on.
The range of possibilities is wide, and one can produce quite entertaining or impressive results.
Leaf skeletons are such beautiful objects that it is strange how few people make a hobby of producing them. They make good subjects for scanning or photography, whether for technical or aesthetic purposes. They are of importance to botanists too, both for the study of physiology, and for identification of species. I happen not to have any of my own pictures to hand, but this one I downloaded from www.vectorstock.com as an example of how attractive they can be.
There are many ways to produce leaf skeletons, ranging from boiling or rotting leaves, to treating them with corrosive liquids. No one method works well for all leaves, and some are very tedious, and take weeks or months, but I here describe one that is quick, cheap and easy, and has worked excellently for me, for many classes of leaves.
It is however VERY DANGEROUS, so follow the safety precautions carefully!
Begin with easy subjects; you can get fancy later.
Look for mature, sound, attractive, thin, flat, firm leaves without blemishes: the older the better, as a rule. Leaves with early autumn colours often are good for most purposes. Leaves from trees such as oaks, planes, camellias, Eucalyptus, maple, fiddlewood, Bauhinia or apples generally are good. Succulent or young leaves tend to end up as mush, though a few succulent species do have usable, elegant three dimensional skeletons when treated by techniques other than what I describe here.
First read the whole description to see what you will need.
Get suitable safety kit. Above all, wear eye protection; spectacles alone are not enough! Even a droplet splashing in from the edge of the spectacles can leave you in need of a corneal transplant, so wear proper comfortable goggles, or better, a full transparent face shield. Some of the cheap face shields on sale for protection from CoViD infection are just the thing.
Rubber or plastic gloves are a good idea too, but not as important as eye protection. A good barrier cream on your hands might be enough if you work intelligently.
Sodium hydroxide (NaOH, "caustic soda" or "lye", not washing soda, which is sodium carbonate) commonly is available from plumbing suppliers as a commercial drain cleaner. It is usually in the form of white flakes. You (or someone) will need to prepare a 10% solution. KOH, "caustic potash" may be even better, but is harder to obtain, more expensive, and even more dangerous, if anything. It has the advantage though, that once you have neutralised your waste, you can bury it because it is a valuable fertiliser. Still, I recommend sodium hydroxide for anyone who needs to ask. Some people use washing soda to avoid working with dangerous caustic soda, but that requires hours of leaf boiling rather the few minutes in which sodium hydroxide works, and on many kinds of leaf, sodium carbonate does not work at all.
As a safety precaution, before you start handling any dangerous soda, get some white vinegar handy and prepare some of it diluted with tap water. You might prefer to use citric acid solution, which is not so smelly; suit yourself as long as your liquid is safe, mildly acid, and immediately available at need. Keep a cloth handy, with which you can wipe up spills or splashes on your skin or clothes. Prepare in advance; don’t do anything creative in a hurry after the accident; a little water fast is better than a lot of vinegar later, but if you have it ready first, dilute vinegar is better than water for dealing with soda on skin or clothing. It also is the right stuff for wiping splashes on the table where it is likely to contaminate your elbows or utensils.
If you don’t have lab equipment, plastic kitchenware will do for preparing the solution. Keep your caustic soda dry and sealed at all times except when actually weighing it out. And having weighed a batch, seal your container at once if you don't want it to cake unusably. All this is partly for safety, and partly because NaOH is hygroscopic and, when left open, reacts with carbon dioxide and humidity, so that it soon degrades to a solid mass of partly-weakened sodium carbonate.
And remember that a good splash of boiling water does not need to contain any caustics to put you in hospital. These procedures are not for the kids, and definitely not for people unfamiliar with good lab practice!
If you are sharing premises with the sensitive of snout, try to keep things family-friendly; remember that leaves boiling in lye have a nasty, soapy smell, so you might want to work in a fume cupboard.
Suppose you will be working in a flattish plastic tray that can comfortably hold about a litre of water. Then put about half a litre in the container in which you will be boiling the leaves (not the plastic tray, but a suitable glass or steel pot. (For goodness sake DO NOT use an aluminium pot!!! Strong caustic soda or potash will destroy it in minutes! The best vessel is heat-resistant glass with a glass lid.)
Gradually add about 100 grammes of the soda flakes to half a litre of water while stirring with a plastic cooking spoon. Add the soda to the water; not the other way round, or things can get hot and messy. Keep stirring till it is all dissolved. Then add the rest of the water. Stir well.
How you boil the solution hardly matters, depending on your equipment; flame, stove-top, or microwave will all do as long as it is safe and you can afford to risk ruining badly chosen equipment.
Heat the solution to nearly boiling. Sodium hydroxide is treacherous stuff; it is inclined to “bump” instead of boiling evenly, splashing everything with a caustic solution, so it is good to have a vessel with a loosely-fitting lid, But once you add the leaves, they generally prevent bumping.
Now add the leaves to the hot water. Don’t pile them in; begin with a dozen or so smallish leaves, such as oak or camellia, or one or two larger leaves, such as some Ficus species, will do well. Use your head, and avoid making a mess. Once the leaves are in the hot water, and not floating on top, boil them for 5 to 15 minutes, depending on how resistant they are. Best start with a short boil and test a few to see how soft they have become. It is easier to return them to the boiling liquid than to unboil a ruined leaf.
Leaves are done when the soft tissues are easy to remove with a soft brush. Put them into tap water in the plastic tray, to work on them. Some leaves, such as those of fiddlewood, Citharexylum quadrangulare, have a couple of layers of membranes and venation, so you might want to keep a pair of forceps handy to deal with the fiddly bits. Other kinds of leaves are just skeleton and pulp. Use your judgement.
Do everything gently. Spread the leaf on a firm spongy surface like newspaper, thin cloth, or something softly rubbery, on the floor of the dish, in the water. The perfect brush should have softish, short bristles. Some of the broad brushes that come free with cosmetic powders are perfect. Usually their bristles are a bit too long, but either you can hold the brush low down, pinching the bristles, or you can guillotine the bristles to suit your personal technique. Toothbrushes and similar devices are unsuitable: stiff bristles damage the leaf veins. Don’t scrape the leaves: hold the brush as if you are gently stabbing the leaf surface with the bristles; keep dabbing to push the pulp from between the leaf veins and leave the skeleton clean. You can rinse each leaf from time to time to see whether you have cleaned it properly.
Rinse each skeleton in water as you finish prodding it clean, then put it into weak vinegar or other acid till you have finished the batch.
This is a good time for bleaching or staining the skeletons: transfer them into weakish peroxide, or hypochlorite, or possibly sulphite, or a suitable dye.
Spread the lot onto a suitable surface to dry; flattening them between clean newspaper or blotting paper works well. This is important, because most leaf skeletons are brittle and easily damaged whether wet or dry, and they will not dry into a good shape unless pressed into shape while still wet.
As a rule one wants them flat, but sometimes one might want to mount them on a curved surface, such as on a glass vase or a lampshade, and then the right time and place to shape them is while they are still wet and soft, and on the right shape of surface. You might want to soak them in a suitable dilute emulsion such as PVA, or a weak plastic solution, to increase their toughness, but that is not usual.
Resin or balsam film for arranging slide subjects.
As a rule it is easy to arrange an object on a permanent microscope slide, usually in Canada balsam under a cover slip, because its exact orientation and position are non-critical. However, when the position really is important, such as when there is a label identifying multiple objects, say an insect’s legs, or even a single object in a given position, then liquid balsam in a xylene solution can be a real pain: you carefully arrange the object in a drop of balsam, and either it moves as you place the cover slip, or, if you wait for the balsam to stiffen you wind up with bubbles under the slide . . . Start over and it happens again — and again. . .
One approach is to create a dam of balsam in a ring of sufficient depth, but that not only looks bad, but causes other difficulties, plus being a confession of inadequate technique. Another approach is simply bung on the cover slip in the hope that some mounts will come out right, and who cares anyway. Well, no one cares of course, except those who are not slovenly, and who cares for them?
I do.
An easy way to get a pretty good result is to pay attention to what happens when you do it wrongly. The typical beginner places his specimens in the balsam on the slide, then places a drop of balsam on the cover slip, then gently lowers the slip from one side, like a hinged lid. Everything works well, till the upper and lower balsam masses meet, then surface tension takes charge, and the whole lot sucks up into a jumble under one edge of the cover slip, along with one’s frustrated self-esteem. A better way — not perfect, but good enough for most purposes, is to lower the cover slip carefully, holding it horizontally with its balsam droplet hanging under its centre, till the drops meet in the middle of the slide. The surface tension sweeps everything up as before, but this time symmetrically. Immediately drop the slip, still horizontally, and the liquid flows back, bringing the objects close to where they started from. It might improve matters to fiddle lightly with the slide for little adjustments, but as a rule, the less you do, the better.
Leave the slide to dry normally, and it is the envy of the uninitiated.
For actual perfection, prime the slide in advance. Put a nice droplet of balsam in the middle of each slide and heat a batch of the slides in an oven just hot enough to make the balsam lose its solvent (usually xylene) and melt into a flattened circular patch that rapidly sets hard on cooling. It must not boil the droplet, because that leaves bubbles that are hard to remove.
Keep as many of these slides as you need where they won’t gather dust. They will last until you want to use them.
When you need to use them, prepare one or two cover slips with the necessary amount of liquid balsam for immediate use.
To set your tricky objects, put them down on the stiff balsam on the slide, and position them as required with forceps and needle with as little balsam as you can manage conveniently. They will stick to the hardened layer of balsam. Add just enough liquid balsam to cover them properly without bubbles, then quickly, but not hastily, while the under layer of balsam is still solid, or at least viscous, place a prepared cover slip horizontally on top. The fixed balsam will hold everything in place for several seconds, and that is how the arrangements will be preserved.
It doesn’t take much practice to get everything practically perfect practically every time.
Dye contrasts.
As I mentioned, I once did some research on aerial spraying. This involved detecting where which kinds of sprayed fluids travelled whither, and how large the droplets were. There were two basic ways of doing this: either dissolve some dye in the spray and catch up the droplets on flat, absorbent surfaces that could be examined back in the lab, or put some dye or dyes on the test surfaces, use unstained spray liquid, and measure the sizes of the areas where the droplets washed out the dye.
The first thing was to see how large an area a given size of droplet covered. This turned out to be more challenging than one might think. We did wind up with satisfactory figures, but only trusted them when we got consistent results when we compared different methods. Ultimately the most repeatable results were when we drew fine capillary glass tubes and measured their dimensions under microscopes, then drew in tiny volumes of liquid and measured their length in the tubes. This enabled us to calibrate droplet sizes against the circular spots they made on given surfaces like art paper or compressed magnesium silicate powder (talc).
The magnesium silicate gave the most beautiful and precise results, but pressing the sheets under hydraulic pressure was expensive and slow, and the pressed slabs were fragile. In the end, art paper, though less precise, was adequate for our needs, even though it meant using dye in the spray.
We could get by without that without staining
the solution, by mixing a solution of two dyes, a water- and ethanol-soluble
green dye (so-called “apple green”) plus an oil-soluble red dye (one of the Waxoline Red dyes worked for me) with the magnesium silicate powder before pressing it. The red and
green were as near as we could get to colour-opposites, and we had to
experiment to get an even grey, because the red was more intense than the
green. One of their necessary attributes was that they did not dye the magnesium silicate, so that solvents instantly left it white where they had washed out the dyes.
The effect was marvellous. Seen under the microscope, a droplet of any oily liquid leached out the red dye, leaving a green disk surrounded by a dark red circle against the grey background, whereas a water droplet left a red disk with a dark green circle. Under green light the green markings literally vanished, while the red dye looked practically black. Under red light the opposite happened. This meant that counting and measuring was much simplified without interference. Alcoholic fluids that we tested out of curiosity, leached out both dyes, leaving dark circles around white patches in white light.
I mention the effect in case it might be useful for anyone working where such discrimination might be useful. As it turned out, we ultimately did not need such fine discrimination, but the effect was interesting and impressive.
Glass spray nozzles
Few of us think much about spray technology, which perhaps is just as well; it is more complex and varied than most people realise. It varies with purpose, materials, infrastructure and needs, just for a start. Just in selecting the nozzles, the uninitiated find themselves in a jungle of rival vendors and designs.
While working on aerial spraying, I happened on a principle that seems to me of some interest and I was almost sorry when I had to let it go, in favour of commercially established equipment that wasn’t especially satisfactory anyway.
The accompanying diagram may help.
The idea is that, without any special constrictions that are easily clogged, input liquid under adequate pressure can be split into two streams. It they are directed to intersect symmetrically, they will form a neat fan of water that sprays nicely.
It behaves very well, and many of its aspects can be tuned by trial and error, such as having the streams intersect slightly asymmetrically for special effects, but for my purposes a nice, symmetrical fan was adequate for as long as I worked on it.
My model I made of commercial thick-walled glass capillary tubing with an internal diameter of a few millimetres. That was fine for internal laboratory applications where there were no labourers to break the equipment, and for prototyping to master the principles. It did not require any special glass-blowing skills, but if you can’t manage the basics, don’t bother.
If you can do the simple glasswork, then first make the general shape and cut the ends to match. No fancy shaping of the ends required. Then heat one of the elbows just enough to permit movement of the tip. Adjust the position of that tip while blowing air through from the base. You can tell when the two air streams match by their behaviour in the flame.
Allow plenty of annealing to relax the stresses from the bending.
Have fun, but, for the sake of your eyes, wear your glasses!
Glass beams for pointers and balances.
While you are in the mood for glasswork, if you are caught short of high quality equipment, consider a home-made micro balance. The power of commercially available modern balances is nothing short of stunning, but they can make a serious hole in a private pocket. I won’t give you a recipe for making your own, but with a few hints you can make quite useful items from resources little more than one might find in quite modest laboratories. One needs a cabinet to keep out air currents because a minute draught can ruin a reading, A discarded transparent plastic container is a good example of the sort of thing you may start with.
The beams of your balance can be made of glass; the main problem is to make them stiff enough: the usual commercial cylindrical glass rods are much too floppy when drawn thin enough for micro-balances. Tubes of relatively wide internal diameter drawn down to acceptable thinness are better, but what I have used most as material for making light glass beams was window glass. Cut waste window glass into strips a few centimetres wide and of convenient length. Heat them cautiously, because they tend to fly apart if you heat them suddenly. Then draw them into flat beams while they are not too floppy, or they lose their flatness. I suppose you could manage some sort of I-beam or T beam, but I never bothered; for the lengths and applications I needed, flat beams resting on their edges were stiff enough.
Professional scales tend to have either fabulous ceramic bearings, or no moving parts, or magnetic bearings. There is a lot of scope for magnetic bearings nowadays with cheap super-magnets commonly available, but you can get pretty good results with round glass on glass if you minimise the balance’s movement. One way to manage this is to give your flat beam a very slight kink at its pivot point, so that the arms bend down enough to prevent its sliding or tumbling off its support.
The next essential is to form weights. For microscale purposes, glass weights are good too. Draw yourself some nice even rods of glass of less than 1 mm diameter, and stiff enough to handle with tweezers. They will look like threads, but I say rods, because you want something stiff enough to handle. Wangle a visit to some lab where they will let you weigh some of your rods so that you know their weight for a given length. Coloured glass is good for easy handling and coding if you can get it.
Now you can bend rods of given weights into hairpin shapes so that you can use them as riders on the scale arms. You also can give them a little bend that makes it easy to pick them up with tweezers. You don't want to spoil them by handling them with fingers. If you have a 1 mg rider weight balancing a load ten times as far from the pivot, that load is 100 microgrammes.
This implies that you need a distance scale that enables you to tell distances along the beam. Fasten a couple of cheap rulers so that you can read the distances without parallax.
As a pointer, attach a fine beam vertically pointing down to a central mark that tells you when all is in balance. The pointer is not for giving a reading, just for telling you when you can take one.
Such a balance can give you readings adequate for weighing ant-sized insects. I have weighed sizeable batches of ants one at a time by mounting them cumulatively, and then calculating the individual weights from the successive differences.
You can have fun even if you don’t have money.
Dead man’s switches & other precautions.
Especially for working with electricity, but also with hot objects or other items that might cause trouble if you panic, get trapped, burnt, or poisoned, it is a good idea before you begin, not after the eggs have hit the fan, to ask yourself what you would have wished you had done if things went wrong. Don’t pour dangerous liquids or pipe dangerous gases that will fall or go on flowing if you get some in your eye or faint or go to sleep or get absent minded. Use sprung or weighted valves, so that if you let go, it shuts off. Electric switches might include a foot switch so that if you let go then everything stops when you move your foot or fall over or go into convulsions. There are many such examples, but you often need to think of them before you find out why they are called dead man’s switches.
It might be a good idea to have some sort of timer alarm when you have a process that takes longer than it is easy or convenient to give sustained attention. Horror stories abound, especially in industrial settings such as explosive factories, when someone has been too clever, but not clever enough to think out the implications of his cleverness. People have died because of someone falling asleep while supervising a process that needed to be controlled. Sometimes an entire neighbourhood has been shattered by the results.
On a smaller scale carelessness in any chemical laboratory can cause premature greying of hair. One item that I retail second hand was when an acquaintance was, for reasons that escape me, preparing some nitrogen trichloride by bubbling a gas through a liquid, with the liquid product floating to the surface whence it was to be collected. Now, nitrogen trichloride is a liquid explosive so sensitive, reactive, and treacherous that sane people do not handle it in more than gram quantities, but this genius went off and forgot about it. Some time later someone else came in and saw the bubbling proceeding merrily with about an inch of nitrogen trichloride floating on the surface of the liquid. Enough to wreck the lab and blind or kill anyone entering at the wrong moment. After an incredulous double-take, he tiptoed in, turned off the gas supply, tiptoed out again, closing the door gently, locking it, and putting a warning notice. They simply abandoned that lab till the nitrogen trichloride had evaporated or decomposed.
I myself am absent minded; in my student days I lived on a farm where my parents had given me an unused chamber for a laboratory, thereby reducing the risks to others' lives and welfare. On one occasion I was, for reasons that now escape me, preparing picric acid in a flask in a water bath on an electric hot plate. It involved a mix of nitric and sulphuric acids, and the reaction took perhaps an hour or so. So I went off for a read. All engrossed, I next thought of the flask when an alarmed employee called me because things were going badly pear-shaped. I ran up to find the room completely impenetrably filled with fumes of the acids and what not. Totally, but totally, unbreathable and uninhabitable. I sneaked round and put an arm into the fog to cut the power to the stove; I opened the outside door to let the room clear.
The water bath had boiled dry and the reaction mix had overheated, squirting a plastic, foaming mass against the ceiling, where it solidified into something like a hemispherical hornet's nest that I kept there for a memento for some months, till someone got into the lab to tidy up, and to my intense annoyance, cleared the ceiling. The fog had included a good deal of picric acid, and had deposited it evenly over everything, including the cement floor. Picric is a yellow substance used for dyeing wool among other things. We had a biscuit coloured great Dane who liked to sleep in the coolth of the lab, and for months afterwards we could see the yellow of his flanks when he had gotten in and slept there.
Well, the lesson was cheap at the price. . .
Drosophila Gem Squash.
When I was studying introductory Genetics before the big luser lost daddy’s first million, we bred our Drosophila melanogaster in the milk bottles that then used to be standard; so standard that one simply did not think of any alternative for breeding Drosophila, except occasionally in Petri dishes for close examination.
Well, I did not become a geneticist, but I did have occasion later in life, to raise chameleons, and the babies of our species of chameleon were too small for a diet of houseflies. They could do pretty well on aphids that were not of any of the poisonous species, but only for a few days, because even inoffensive greenfly are not a balanced diet, and even greenfly produce a bit of the Hemipteran repugnatorial stinkbug smell, though most people don’t realise it; and that is harmful for tiny reptiles. Drosophila were far better baby food, but at first a bit harder to organise.
I experimented with several cycles of production trials, including borrowing some milk bottles from the genetics department. It worked, but the loan was a bit grudging, because it turned out that glass milk bottles had since gone out of fashion, so that what had once been a cultural fixture, was now unobtainable for love or money. I was however not in need of nice controlled cultures, just cheap, disposable media for mass production of flies.
What worked was to choose some kind of vegetable shell that could stand a judicious bit of cooking without coming apart. Scooping out the inside of a thick-shelled citrus, or a large granadilla did well. My favourite however, is the shell of an overripe gem squash: a spherical vegetable that can be eaten green or ripe, but is best when eaten peel and all when unripe. However, when fully ripe, its shell is almost wooden, and that is the stage I refer to in this description.
Take such a fruit, in particular a ripe, hard gem squash, cook it till the inside is soft, cut off one end to leave a hole of a size suited to a teaspoon, and scoop out the unwanted part of its insides to be disposed of as desired, then half fill it with Drosophila medium. In the case of the squash, if the pulp of the fruit is not required for human consumption, one need only remove the seeds and cook the gourd till its pulp is tender enough to stir in the rest of the medium. Judicious microwaving works well, but so does (equally judicious) boiling.
With or without gem squash pulp, one prepares a medium that is something digestible and porridgy. Cooked oatmeal, on the sloppy side, with a bit of milk and minerals such as a bit of dolomite powder and some sugar, worked for me. About half of the volume of the shell is about the right ration for a gourd. Let the mix cool down to about blood heat and not more than 40 degrees C. Cooler if in doubt. Stir in yeast enough to get it fizzing within an hour or two, and leave it where Drosophila will find it within a day or so in any warm season. Anywhere near garbage or a greengrocer’s shop will usually do nicely, but if you are in a rural area or wherever anyone else is breeding Drosophila, they will find you easily enough.
Any yeast that will set the mix fizzing will do. If it does not froth within a few hours, then add more yeast. Bread yeast from shops commonly is the most convenient, but fruit peels or sourdough generally are good starters as well. Once fermentation is well under way, you can use some of the cultures at once for Drosophila production. You also can keep a supply of the cultures in the freezer and take them out to restart fermentation again when required. Do NOT freeze insect eggs or larvae; it will kill them.
When you see flies beginning to emerge from the hole in a week or two, just put the culture into the cage or wherever the chameleons can reach them as they emerge, and they will do the rest.
Let the chameleons feed ad lib on the flies visiting or emerging from such a culture for as long as you like; they will come to no harm such as they might from aphids; that is the point of the milk and minerals in the medium; the flies not only are nutritious themselves, but their gut contents will be digested as well.
Other insects will appear too, but only ants are likely to be a problem, and you need to protect your animals from ants anyway. Argentine ants can wipe out a brood of baby chameleons overnight, and that is a horrible experience.
Trust one who has experienced it.
A given culture will last for a few weeks if all goes well, but it depends on conditions. It might be necessary to moisten it with water or juice or milk from time to time. The culture changes as it ages, and different species of flies will emerge, some more welcome than others; I remember some tiny black wingless ones that I suspect must have come in on the backs of visiting Drosophila. They were too small to be useful, so I ditched the culture, but I never saw any sign that they were doing any harm. Studying them might have been rewarding, but it was not convenient at the time.
Exhausted or simply messy cultures can be disposed of to pigs, added to compost, buried, or if you really have no options, dumped into garbage. Just do NOT try to flush them down lavatories, or you will regret it.
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