Search This Blog

Friday, 30 December 2011

Making your own microscope

Robert Hooke's 1665 microscope only
had a single lens

If you do a web search on <USB microscope> (leave out the "angle brackets"), and dig around, you will find instructions for making a simple 'microscope' from a webcam that plugs into a computer. Be warned that you need to be fairly handy with tools and a soldering iron to try this sort of thing. You might be better off looking around for one of the cheap models that is available, like the one I showed a while back.

Remember that good work can be done with just a single lens. Anton van Leeuwenhoek did amazing work in the 1600s with single lenses, and the tradition continues. Try a web search on <water drop microscope> and discover some neat designs for one-lens microscopes. Or read this, which appeared first in Charles Dickens' Household Words and then was reprinted in Scientific American, November 4, 1854, p. 64. Maybe you can see how to make such a "microscope" yourself.
 There is a man who sometimes stands in Leicester square, London, who sells microscopes at one penny each. They are made of a common pill-box; the bottom taken out, and a piece of window glass substituted; a small hole is bored in the lid, and therein is placed, a lens, the whole apparatus being painted black.
Upon looking through one of these microscopes, I was surprised to find hundreds of creatures, apparently the size of earthworms, swimming about in all directions yet on the object glass nothing could be seen but the small speck of flour and water, conveyed there on the end of a lucifer match, from a common inkstand, which was nearly full of this vivified paste.
I bought several of these microscopes, determined to find out how all this could be done for a penny. An eminent microscopist examined them, and found that the magnifying power was 20 diameter. The cost of a lens made of glass of such power would be from 3s. to 4s. how, then, could the whole apparatus be made for a penny?
A penknife revealed the mystery. The pill-box was cut in two, and then it appeared that the lens was made of Canada balsam, a transparent gum. The balsam had been very cleverly dropped into the eye-hole of the pill-box. It then assumed the proper size and transparency of a well-ground lens. Our ingenious lens maker informed me that he had been selling these microscopes for fifteen years, and that he and his family conjointly made them. One child cut the pillbox, another the cap, another put them together, his wife painted them black, and he made the lens.
It all sounds too easy, doesn't it? I suspect the man in Leicester Square and his family may have had a great deal of practice!

Coming up early next year: looking at a feather and comparing sand samples from different sources. I have been distracted because I needed to buy an Android tablet for my writing work, and I have started playing with it in a serious way.  The writerly side of me may look at the many meanings of table and tablet at some stage.

Thursday, 22 December 2011

Closed for Christmas

That's it.  I'll be back around December 28.

I'll be off out in the bush, looking for beasties like this.

Go on, get away from the computer and get outside as well!

And seasonal greetings for whatever season you are currently choosing to celebrate.

Tuesday, 20 December 2011

A plankton net for collecting small animals

You can catch lots of interesting small water life by dragging a bucket on a rope though water weeds and then filtering the results.  A piece of ordinary denim makes an excellent filter, as you can learn if you Google <copepods denim cholera>.  Once you have filtered the water, wash your "catch" into a clear container and hold it up to the light.  use an eye dropper, a Pasteur pipette or a modified wash bottle to extract animals from the sample.

If there are no water weeds, you will need a net.  Even a home-made plankton net can sample small animals at or near the surface of the water.

The main parts are a towing line to pull the net along, a swivel at the net end of the line to stop the line kinking as the net spins while it is being pulled, a stiff hoop to hold the net open, a very fine net (think about what you have in the scraps basket at home), three lines attaching the hoop to the swivel and a small glass bottle which attaches to the lower end of the net, a metre or so beyond the towing ring.

You can buy a swivel from any fishing equipment shop. The hoop can be a length of coat hanger wire, bent in a circle, the mesh can be the footless leg of a stocking, either stitched or stapled over the hoop, or even glued to the hoop with contact adhesive (wear gloves and work outdoors to avoid the fumes if you use this glue). A stapled net won't last as long, but it's easier to make.

As you pull the net along, any animals trapped in the open mouth will be pushed down to the end, where they will be largely protected from damage by the still water in the glass bottle. You can buy nets like this for a high price, or you can make your own very cheaply.

The towing line can be a fishing line: I have used a fishing rod to haul a net like this through the water while walking along a wharf or jetty (the rod stops the net from snagging on the pilings), or you can tow one from a boat which is being pulled along by a 2 hp motor.

If possible, fix the bottle to the net by attaching a metal or plastic screw lid at the narrow end of the net, so you can change bottles regularly, just by unscrewing them. That lid will need a hole in it, but if you use a standard jar, you can have plenty of spare lids. Otherwise, tie string around the stocking and the bottle, and pull it very tight.

With a suitable net, you can explore the plankton types and densities over a time period: either looking for daily patterns, or monthly patterns (some plankton may respond to the full moon, so samples taken regularly at 9 pm could be useful). Maybe there are patterns you can see as the seasons change.

Nets like this can also be hauled through seaweed and water weed to sample the small animals living on those plants. This is likely to damage the net, so use a replaceable but strong one.

You could also just explore the types of plankton found in one place, or compare different environments at more or less the same time of day, over a period of time, to see whether any observed differences continue over long periods. Aside from that, you have the tool, you have some ways of using it, so go for it, remembering that the most interesting questions are always your own questions!

The origins of the towing net

Nobody knows now who was the first to develop this handy item. John Macgillivray, writing in the 1850s, thought it worth explaining how one was made, so maybe the idea was new back then:
Not having seen a description of this useful instrument, I may mention that the kind used by Mr. Huxley and myself, consisted of a bag of bunting (used for flags) two feet deep, the mouth of which is sewn round a wooden hoop fourteen inches in diameter; three pieces of cord, a foot and a half long, are secured to the hoop at equal intervals and have their ends tied together. When in use the net is towed astern, clear of the ship's wake, by a stout cord secured to one of the quarter-boats or held in the hand. The scope of line required is regulated by the speed of the vessel at the time, and the amount of strain caused by the partially submerged net.
—John Macgillivray, Narrative of the Voyage of H. M. S. Rattlesnake, vol. 1, chapter 1.
Or maybe it was only new to Macgillivray.  As early as 1768, Joseph Banks makes mention of using both a "cast net", and when that was lost overboard, he attached a hoop net to a fishing rod. Perhaps this was just dipped into the water, but the idea of towing a net seems obvious enough. Like a lot of simple ideas, most people probably thought it not worth mentioning or explaining!

Sunday, 18 December 2011

Slowing small water animals down

Well, as promised, here are some notes on getting to actually see live animals in a well slide like the one on the right. This is a standard glass microscope slide, 3" x 1" (75 x 25 mm), but with a small depression cut into it, so that a cover slip can lie flat on the slide, even when a large-ish (1 mm or so) animal is there is a wet mount.

The problem is that live animals swim around and go out of the field. They also go up and down and go out of focus.  That means you need to slow them down.

The three main ways of slowing animals down are:

* to put barriers in the way, so the animal can still move as fast, but not as far;

* putting the animal in a more viscous (sticky) solution which usually kills them in the end; or

* kill them outright.

The most common barriers are bits and pieces of cotton wool or ground-up face tissues. This is not very effective with anything smaller than a mosquito wriggler, but it's better than nothing.

Live specimens can be mounted successfully in ®Gurr's Water Mounting Medium, which slows them down (and kills them). I have been using the same bottle of this product for almost 40 years, and it seems to be hard to buy nowadays, though it is still mentioned by professional scientists.

A solution of 10 g of methyl cellulose in 90 mL water forms a syrup that will slow most animals down for microscopic examination, while allowing observation of movements of the gut, breathing tubes, and so on. You can buy methyl cellulose at hardware shops, though you may also get it at health food shops, where you will probably pay a lot more for it. Be careful not to get it in your eyes or on your skin.

I haven't tried this, but I'm told you can also add 2-3 grams of gelatin to 100 mL of cold water and heat this while stirring. Cool the gelatin solution back to room temperature and add one drop of pond water to one drop of gelatin solution.

If you mount the animals in 70% alcohol, this will kill them, but a 1% solution of magnesium sulfate (often sold as "Epsom salts") will just anaesthetise them. Note that 1% here means 1% by weight or one gram in 100 mL of water. Put a drop of this on the slide and then use a camel hair brush to add the animal.

Just a reminder for those coming in late: the material I am posting here is made up of out-takes from an upcoming book Australian Backyard Naturalist, due out in May 2012. This is the stuff that won't be there.

Next time, I will look at catching nematode worms.

Saturday, 17 December 2011

Small water animals

This entry is about the small crustaceans we call water fleas, because they are about the size of fleas, and they live in water.

Water fleas at a glance

These animals may be flea-sized, but they are actually crustaceans, distant relatives of crabs, prawns and slaters. The ones you are most likely to see are Daphnia, Cypris and Cyclops, but you never know your luck! They move differently, but you need at least a hand lens to see any details, and they are excellent for low-power microscopy.

Technically, they are all branchiopods (not to be confused with brachiopods!). Daphnia are in the sub-order Cladocera, the similar looking Cypris is in the Ostracoda, and Cyclops is in the Copepoda, so you may need to look up cladocerans, ostracods and copepods to find them on the web. The copepods are much less flea-like.

Branchiopds are easy to collect, because they will be found in most bodies of water, and they are just as easy to cultivate. They also have some interesting biology: the Cyclops that you see here is carrying two egg sacs, and you can often see eggs inside Daphnia.

I want to begin, though, with an odd discovery about branchiopods. It was made by Jacques Loeb, a German-born physiologist who moved to America. Loeb made some important discoveries on how animals respond to stimuli, and also did some useful work in embryology. He never explained how he made this discovery, but it must surely have been during a laboratory party!
The writer found that certain freshwater crustaceans, namely Californian species of Daphnia, copepods, and Gammarus when indifferent to light can be made intensely positively heliotropic by adding some acid to the fresh water, especially the weak acid CO2. When carbonated water (or beer) to the extent of about 5 c.c. or 10 c.c. is slowly and carefully added to 50 c.c. of fresh water containing these Daphnia, the animals will become intensely positive and will collect in a dense cluster on the window side of the dish. Stronger acids act in the same way but the animals are likely to die quickly. . . Alcohols act in the same way. In the case of Gammarus the positive heliotropism lasts only a few seconds, while in Daphnia it lasts from 10 to 50 minutes and can be renewed by the further careful addition of some CO2.
— Jacques Loeb, Forced Movements, Tropisms, & Animal Conduct, Dover edition of 1973, pp. 113–114.
In the passage above, 'heliotropism' means "moving towards the sun". People now prefer to say 'phototropism', meaning "moving towards the light", instead. Strictly, heliotropism means "moving towards or away from the light", which is why Loeb speaks of a "positive heliotropism” to show that the animals moved towards the light. Negative heliotropism would involve a movement away from the sun.

Today, we can see the logic of the animals' reaction: high CO2 means less oxygen, so moving towards the light usually means moving upwards and getting closer to the oxygen-rich surface layers of the water.

As a rule, when you are cultivating water animals in bottles, leave the water level far enough down to keep the surface area large. This maintains oxygen levels.  On the other hand, if you want to collect animals to look at, fill the bottle almost to the top, and within 24 hours, most of the small crustaceans will be in the top centimetre or so.

The tiny crustaceans (which is what they are) thrive wherever there is food, so green water from a pond will usually have some, but puddles, horse troughs (if they still have those where you live) and so on are also worth trying. Now for the rest of this, I am going to call them all Daphnia. At its simplest level, half-fill a bottle with green water, add a pinch of all-purpose fertiliser, cover it to stop mosquitoes getting in or water flowing out too messily if it tips over, and leave the bottle in the sun for a week or so.

The best bottles to use for this are 2-litre (or larger) PET plastic fruit-juice bottles, with the labels scrubbed off.  PET plastic is clear, so you will be able to see the animals if they are there. They show up best when you stand the bottle on a table in sunlight, crouch down and look towards the sun, especially near the top of the water and loom for small dots that are moving around near the surface.

As a general rule, that is all you need to do.  On the other hand, some professional biologists prefer to feed their Daphnia on small amounts of brewer's yeast, so the choice is yours. The golden rule is to have several cultures of anything precious, and to feed them at different times. That way, if the yeast takes over, you will have other cultures to fall back on, though usually, if a 'dead' culture is left for a while, there will be eggs, spores or survivors which will bounce back.

The best way to breed large numbers of Daphnia quickly is to take some water from a murky green aquarium, without any filamentous algae. Add a small amount of hard-boiled egg yolk, mixed with water into a sort of soup, and stand back! Any Daphnia that you picked up with the algae will start to breed very rapidly.

In stagnant water, Daphnia develop more haemoglobin, up to ten times as much as in water with plenty of oxygen, so the Daphnia from stagnant water can be quite pink in colour. See if you can observe this.

To look at these animals under the microscope, you need well slides, and you need some method of slowing the animals down, so they don't whizz out of sight.  I will talk about that next time.

Wednesday, 14 December 2011

Stains in microscopy

This is probably the most technical and difficult section in this blog (and also the least illustrated). If you are not planning to cut thin sections and stain them, skip over this.  If you persevere, pay close attention to the safety messages.

Most of the things inside a cell are transparent, so it is hard to see any detail. A dye that attaches to one kind of cell part makes it show up more clearly, and we call that dye a stain.

There is a catch, because dyes often cause difficulties with authorities like parents, landlords (or landladies) and the like. These stains can easily stain baths, basins, carpets. people and pets—among other things. More importantly, while pale and colourless chemicals can also be dangerous, you are usually wise to assume that coloured chemicals are always dangerous. Any chemical which attaches to a biological molecule inside a cell (as stains do) is likely to cause damage in the cells that are attacked. Treat all biological stains as dangerous, to be on the safe side.

That means you need to discover the MSDS, the Material Safety Data Sheet. These are easy to find on the web by searching on <(name of chemical) MSDS>. Some MSDS sheets are hard to understand, but the ones at are reliable and clear. Try searching <methyl cellulose MSDS> for practice.

We are forever learning new things, and while methylene blue is regarded as safe now, that may change. Read the MSDS first, before buying or using any stain! You also need to understand that an MSDS will spell out all the risks: read the MSDS sheets for table sugar (sucrose to chemists), water and table salt (sodium chloride), and you will see how complete and obsessively thorough they are!

To get more information on the web, I suggest a search such as <microscopy stains safe>. Just be careful about what you believe!!

Here are some stains that I regard as fairly safe. Even so, you should handle and mix any stains out of doors in good weather. If the stain comes as a powder, think safety first. When you take the lid off, there is sometimes a puff of dust that you don't want to breathe. Try to choose a day when there is no breeze: if there is a light breeze, stand upwind of the bottle. Never mix stains in high winds. Use gloves, goggles and a face mask, or if possible, buy the stains as solutions.

In this outline, I indicate the uses for which each stain is most often used, but most stains will work on other tissues as well.

Basic fuchsin: used to stain nuclei. Dissolve 0.1 gram of the powder in 150 mL of distilled water and add 1 mL of 70% ethanol.

Eosin Y: used to stain muscle fibres, cytoplasm and collagen. Dissolve 1 gram in 100 mL of tap water.

Methylene blue: used to stain living organisms. Dissolve 1 gram in 100 mL of distilled water and add 0.5 gram sodium chloride. This stain can be obtained as a solution from some pet shops, but it will need to be diluted and may have nasty additives. Remember that this one will stain sinks, basins, baths and toilets—and skin!

Nigrosine: used to stain bacterial spores and capsules. Dissolve 1 gram in 20 mL of water.

You can also experiment with food colourings. Many of these are now accused of being dangerous, even when they are approved for adding to food. Treat them carefully, just in case.

Iodine: this is not the friendliest of materials, but it's the best stain for starch in plant materials. Add iodine crystals to a saturated solution of potassium iodide in water until it is saturated, filter and dilute to a pale golden brown. Check to see if you can buy 'tincture of iodine' from your pharmacist, but it will be expensive, and these days, in Australia at least, you will probably only be able to get it from a specialist pharmacist called a compounding chemist.

Malachite green: is not so safe but it is useful to stain plant cytoplasm. Dissolve 1 gram in 100 mL of tap water. This is available as an anti-fungal solution from aquarium shops, but that solution usually contains formalin, which is really dangerous. Read the label first!

When you come down to it, the staining of thin sections might be Too Much Trouble, so what else can you do?

One easy observation involves the large cells that are found in a layer called the epidermis on a piece of onion.  I touched on that a while back in A bit more about microscopes and hand lenses — and I may get back to it at some stage.

Tuesday, 13 December 2011

Normal service will be recommenced shortly

I am on my way back home now, having been across the Tasman, playing with grandchildren, bashing through the very last check of Australian Backyard Naturalist, sent over the seas as a PDF, and in between times, gathering sand samples for further study for that microscopy project on sand.

Sunday, 4 December 2011

Using a pooter (or inhalator)

Just a quick one today, because I have been doing a lot of heavy data-shovelling for the gold book.

This is an old-style pooter, which you should avoid like the plague, because it is dangerous.

This dangerous and old-fashioned design used a glass jar, glass tubing and cork.  I was helping at a Cub camp one day, and my task was to look after a hyperactive kid and keep him interested for the day.  I showed him how to find small things and catch them with a glass pooter, and he was delighted.

That was fine—in fact it was in the specs I had been set for him to be delighted, but he set off, whooping and hollering, leaping over rocks with this glass jar and I just knew that he was going to fall and gash himself. He didn't, but I knew I had to do better.

A couple of weeks later, I was running a workshop for teachers at the Australian Museum. It was all about using scrap and junk to do real science, and I showed them where I was at. The second and third pictures show the solution,  My thanks to Carrie Bengston, who drew the third pic.

In a flash. one of the teachers suggested using a film canister for the job.  This was almost twenty years ago, and people used 35 mm film that came in canisters.  For years, I would go to my local photography shop and come home with plentiful supplies.

Digital photography killed the photography shop, and my supply dried up.  I needed a new design that used components that would be available for at least the foreseeable future.

Well, you can see me making (1) the film canister version; and (2) a newer version that is featured in Australian Backyard Naturalist, if you go to the video links given in the last paragraph.  There's nothing special about these designs (except that they work),  But it's how they work that counts. There are four absolute requirements:

(1) You must have a clear container, and
(2) You must have a lid that comes off easily, and
(3) There must be a cloth filter to stop wee beasties entering your throat, and
(4) There must be no breakable parts.

The two designs you see in the video meet all of those criteria. Look at the video, then come up with your own design.

The don'ts:

* Don't pooter up ants, because they make formic acid, which burns the throat;

* Don't pooter up stink bugs (think about it);

* Don't pooter up millipedes, because their secretions may be toxic; and

* Don't even try to pooter things that are bigger than the tube (think about that!).

To get some more information on making pooters, you need to look at a video on pooters that I made for the National Library of Australia.  I actually did three videos, which you can find from here.  (By the way, that was me before I found out that I was in reach of being overweight, and also that I had a genetic predisposition to diabetes.  I didn't need to be told twice, so I now cast a far smaller shadow.)

* * * * * * *
This blog covers quite a few different things, so I tag each post. I also blog about history, and I am currently writing a series of books called Not your usual... and the first two have been published by Five Mile Press, The offcuts appear here with the tag Not Your Usual... . For a taste of Australian tall tales, try the tags Speewah or Crooked Mick.   For a miscellany of oddities, try the tag temporary obsessions. And language us covered under the tags Descants and Curiosities, while stuff about small life is under Wee beasties.

Saturday, 3 December 2011

The art of the white dish

Some of the most interesting things to look at are the tiny animals that you will find in any pile of old vegetation, but the trick is to make them show up.  Most small animals are very good at keeping still, because predators are all very good at seeing movement. Prey animals are good at seeing movement, because any moving thing might be a hunter. Our ancestors were probably once both prey and predators, so we have inherited the same ability.

The best way to see life in a rock pool is to sit and watch, but exactly the same method works almost anywhere in nature. Sit still in a tree's shadow on a night with a full moon and you may see possums, bats, birds and more, up in the trees.

Using a white dish makes it even easier to see tiny animals moving. When you pick up some leaves or grass clippings, there may seem to be no life there at all. If you spread the material out on a white surface and wait, you will start to see small animals moving around cautiously, looking for somewhere to hide. You can use white paper for this, but a dish reveals them well and also stops things escaping (except the jumpers!).

You will need either a camel hair brush or some sort of probe (a stick, a piece of wire, an old pen or pencil) to move the litter around. If you do this in strong sunlight or under a bright lamp, any animals you uncover or dislodge will scurry off to the nearest shelter, and you can see the movement.  I use an old white enamel dish, the sort your great-grandmother may have used in the kitchen.

These are heavier than plastic dishes, but they are useful for many things, as you will see above, where it is functioning as a home for ant lions on the left and an algae tank on the right. You can also use the dish, with salt water in it, to shake off the small animals clinging to a piece of seaweed. Fill the dish, swish the seaweed vigorously back and forth in the water, remove it, and look to see what is darting around.

On a side issue, semi-clear 3-litre food boxes make good small tanks. It is easier to see what is in these if you sit the base on a piece of white paper or board.

For land animals, you can turn the dish into a cage, if you use flywire and rubber bands to keep them in the container. Watch out that the contents don't dry out too much.

You can see from this shot how I join rubber bands together and then close the loop with an opened-out paperclip. It is always wise to use two separate sets of bands, just in case one of them fails, and gives the animals a chance to escape, to the annoyance or horror of your family.

Tomorrow, I plan to say something about the pooter (sometimes called an aspirator), and how you can use it.  I won't say a lot, because pooters are still there in  Australian Backyard Naturalist, but there's a video already up on my publisher's website, and I have a bit more to say.

Friday, 2 December 2011

The hidden value of performances.

"Those really were two unforgettable hours.  It's been a long time since I've been able to concentrate so well on my problems with arsenophenylglycine.  We'll have to make a small substitution the first thing tomorrow."

That was, according to an article published in New Scientist 22 August 1985, p. 48, the comment made by Paul Ehrlich (1854 - 1915), to his wife as they left the concert hall.  That was my notes say, but in these modern days, you can Google and see the book review that it actually appeared in.

Lehar's bust in the Stadtpark, Vienna, found  while
idly wandering  through there in 2006. The committed
statue photographer can have a great time in Vienna!
Last Tuesday, I was at the last night (in Sydney) of the Australian Ballet's production of The Merry Widow.  I have seen it before, but a month or two back, I saw the Australian Opera's production of the operetta.  
The handy label, set in the lawn nearby for
foolish and confused foreigners who
assume that in Vienna, every statue is
Sigmund Freud, Johann Strauss or Wolfie

While I had seen Franz Lehar's The Merry Widow before in German (in which I have bare survival skills), it was only then that I "got" the  complicated plot line.  The operetta's plot is rather slimmed down in the ballet, so a lot of the Gilbertian foolishness of the operetta is left out, but John Lanchbery's arrangements of Franz Lehar's music are loads of fun, and I thoroughly enjoyed the evening.

But, and I hesitate to confess this, I entered the hall, where we sat in the same subscriber seats that we have had since the Australian Ballet first performed at the Opera House in 1973, with a puzzle in my mind, and I walked out with a new narrative structure for the next book, which I have been researching for some years, gathering the data.

I store all of my research notes, quotes and snippets in a spreadsheet, on which I later play cute tricks to get an order. You can see some of my methodology here, but the main point is that I do free-form fact gathering, chasing interesting bits, and later, I choose from among them, dig some more where I need to, and not long afterwards, the story takes shape. To give you an example, the next book will be perhaps 60,000 words, maybe less, but there are almost 250,000 words in my files at this stage, in about 2800 lines on the spreadsheet (with other sheets in the spreadsheet storing references, image details and other stuff that I will need.  My rule is to over-research, and then pick out the most interesting bits.

This time, though, I had lots of interesting anecdotes but the narrative flow wasn't there. I had sorted everything, but it just didn't gel. But now it has.  It began with something I read, just before we walked down to the Manly ferry, which I told Chris about as we dined in the foyer. It was an October 5, 1851 report in the Sydney Morning Herald about events at Sofala the previous Sunday.  That, I realised, somewhere in Act 2, was the kick-off point, and everything follows from it.

Sand from a Sydney bush track, x10.
This morning, I have been slapping down headings and heading sequences, and it is going to work. I also have in-principle interest from a publisher.

So now I am prepared to come clean, though if you search previous posts carefully enough, I have referred to the idea before, though I was then contemplating a world history—until I saw how big the story was.  Now it will be an Australian story, with many visits to parallels in other times and places.

Sand from a Sydney bush track, x60.
The subject will be gold and gold rushes, which you might have worked out from Sofala, 1851.  It will be a social, a scientific and a technological history, and it will concentrate mainly on the real costs of gold mining.

But the microscopy won't be stopping.  I am still researching the book after that, and that means I will be looking at a lot of sand.  Does that sound mysterious?  If so, good!

To do that sort of work, I think the cheap microscope is going to prove adequate, but it's summer here, and I'm hitting the beaches with clip-lock bags, taking lots of samples to bring home and study.

I'll let you know in a couple of weeks how useful it was, and which lighting proved to be the most useful.

Thursday, 1 December 2011

My current writing status

Just turning away, briefly, from microscopy, I am emerging from a hectic period and moving into an intense one.  My sequel to Australian Backyard Explorer, which is, like ABE, directed notionally at ages 10 to 14, and called Australian Backyard Naturalist, has gone to the printer.

The book after that is for the general market, and the working title is Curious Minds. The title is a play on my occasional statement that my friends and my enemies agree, with differing intonations, that I have a curious mind.  It is about the natural historians and natural history painters who were in Australia, mainly between 1688 and 1888.  That is now through editing and on its way to design, so that will generate a few calls on my time.

The next book is a toss-up.  I have two interesting topics: one is just about completely researched and planned out, a fascinating mix of Australian history and lots of science and technology.  The other, less researched but involving a lot more field work, is starting to push ahead.  This is why I am approaching an intense period, because between Christmas and New Year, I plan to start laying down the text of the new book, and I need to be in a position to decide.

Mind you, there are also three series that are in the offing.

I may have mentioned that I never get writer's block: I just shift to another project for a week.

So if I fall silent for the first half of December, I am most probably still on my twig, but I'm cogitating.

Looking at moth scales.

Butterfly and moth wings have no colours in them—or at least there are no pigments. The scales on the wings produce the effect of colour because of the way they catch the light and bend it. If you want to look it up, the process is called birefringence. Even though you will probably want the higher magnification of a monocular microscope, you may need to use reflected light to see any detail.

That is, you may want to try using a bright light above the stage, though the shots above were taken with my good monocular microscope, and at the magnifications marked.

Note that magnifications stated like this are a bit meaningless, because then I make a JPEG, resize it, and you view it at some uncontrolled size on your screen.  To have any meaning, what I need is a scale, and I haven't worked out how to do that.

Anyhow, here's what you need, snapped on the desk where I am now working. You need a microscope (out of shot), slides, cover slips, a piece of black paper, microscopy tools, and one or more dead moths and butterflies.

The best place to collect dead moths is in an exterior light fitting that is on at night.

You need then to scrape some off, add water and make a wet mount of them.

This is easy to say, but harder to do, because sometimes the scales repel water and slide out from under the cover slip. In the end, I took a dead moth, snipped a few pieces of its wing and dropped them into a small specimen tube with a few drops of water and a tiny amount of detergent. This is why the first shot features an annoying air bubble in each magnification. Like life, microscopy often involves trade-offs.

Here are three shots of the main parts of the process of making a slide. I added a bit much water, so I then had to blot the excess away with a face tissue. By the way, I used the black paper for contrast, so I could see the scales on the slide. It's a handy trick to remember.

The slide in these shots gives you a scale of sorts, because it is 25 mm (one inch) from top to bottom.

 These three shots were all taken with the $50 USB microscope which will appear later in this entry.

The first one is at x10, the sort of magnification you get from a normal hand lens, and as you can see, it isn't much use.

Now we are at x60, and like the first shot, this is using transmitted light, which means I have turned on the light under the stage.  Now we can actually see some detail.

Let's move up to x200, the maximum that I can get. It's a bit pale, and the focus control on the cheap microscope leaves a lot to be desired, but hey, what do you want for 50 bucks?

 Anyhow, I decided to try oblique lighting from above, using a little quartz-halogen lamp that sits on my desk.

The lower shot is one of the pictures I took, but I don't think I'm really in control of this method yet. Anyhow, play with it!

Now a note about my first way of getting the scales from the moth's wing: I think the method shown here, where I just rubbed the scales off with a pair if forceps worked best,

I used scissors to cut bits from a wing.

I used the handle of a paint brush to treat the wing pieces roughly, before I fished out the main pieces of wing. Then I stirred the water up a bit and lifted a drop of it to make my slide.

There were only a few scales on the slide, and some of those were broken, but there were enough whole scales to study.

One thing I learned: the scales seem to vary in any one moth, but I don't know whether each type makes a different colour. There's an interesting bit of research for somebody there!