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Tuesday, 29 November 2011

Looking at pollen grains


What you need:
The first thing you need is a collection of flowers. I use a set of neat little plastic containers that screw together. They are sitting on my desk as I write this, and the top two contain millipedes that I collected this morning, and which may be the basis of a future entry, depending on what I can see.

You really need a microscope with a magnification of x400 or better, and the patience to piece together different views. Pollen grains are too small to see with the eye, or even with a hand lens, but they are too large for you to focus on the whole grain at one time under high power.

That means you need to manipulate the images with a neat bit of open-source software from the National Institutes of Health in the USA, called ImageJ.  This is really neat, but not entirely intuitive to non-geeks, so I really urge you to read the documentation.

Almost every species has a distinctive pollen, and there is even a science of pollen study called palynology. This science is useful in archaeology, criminal investigations—and even in determining the origins of honey.



Here are two shots of the same pollen, one under low power (left) and one under high power (below).  You need to adjust the focus up and down if you want to see the distinctive patterning (called sculpting) on the surface.





The pollen is from a pea, Gompholobium, which was quite infamous in the early days of Australian settlement, because it poisoned sheep which ate it. The poison, sodium fluoroacetate, is used today under the trade name "1080" (ten-eighty) to get rid of pests, because Australia's marsupials are able to resist the poison which is good against rabbits and foxes.





I didn't get very far with this before deciding to drop it from the book, so the lighting in my examples is quite variable, simply because I had lots of stuff to write, and couldn't chase too far down the blind alleys.  I will get back to this, one day, some time. Take these two shots (below) of cobbler's pegs (Bidens sp.) pollen:



In each case, I took three shots at different levels, trying to capture the sculpting. Here is a last example, Kunzea (bachelor's buttons, Myrtaceae).  I'm far from an expert in this, but there's the idea for you.


There are two problems: pollen grains are sometimes hard to wet, so air bubbles cling to them, and once they are wet, some of them will burst, sending out a pollen tube—this is explained in the next section. I use a tiny amount of detergent in the mounting water and that seems to help get rid of the bubbles. To beat the bursting problem, all you can do is work fast.

Professionally, the clearest views come from scanning electron microscopes, but those are a bit more than a private individual can afford. Under x200, you will be able to just make out the 'sculpting' on the surfaces of pollen grains, and you may even be able to see that they are different shapes. Under x400, it will be much easier to see.





Sunday, 27 November 2011

And now I am on steam wireless—again

Another side of my persona: aside from wandering in vast wildernesses like that on the right, I am an old educator (well that will come as a complete surprise, I don't think, if you have been reading these pages).  I also write books, and I haunt libraries.

Add to that my long history with computers, which began in 1963 when punched paper tape was a modern form of input.

I am also a former bureaucrat and as a management consultant, spent some time doing fraud investigation, so I have a short patience span when it comes to would-be pole climbers who seek to wreck the status quo, so that they can later point proudly to the carnage and say "I created that."

All of these elements come together in a thoughtful piece that I delivered today on ABC Radio National, called A Question of Collaboration.

For a certain period of time, you will be able to listen to it, and after that, you will be able to read it.

I have made many contributions to Ockham's Razor since 1985, and you can find most of them here.

I can't see any way to attach an mp3 file to this blog, but I will play with it, once iTunes has gathered it in as a podcast.  You can subscribe to the podcasts here.

(Later addendum: I looked too soon—the ABC elves have since added a link so people can download the file.  It will be there until about Christmas day.)

I could do worse than quote from the program's own About Us page:


William of Ockham was an English monk, philosopher, theologian and probable victim of the Black Death, who provided the scientific method with its key principle 700 years ago.


'What can be done with fewer assumptions is done in vain with more,' he said. That is, in explaining any phenomenon, we should use no more explanatory concepts than are absolutely necessary.


Well, for both broadcasting and for science, simplicity should never be despised. Our program, named after William, consists of a short introduction followed by a scripted talk. Just that, week after week.


This program allows thoughtful people to have their say without pesky interviewers interrupting, or someone of opposite views turning the exercise into a joust. There are times when a speaker needs a clear run, some proper control, and this is what Ockham's Razor provides. 


Have a listen or a read, and see what you think.  You need to get to the very end.  In the picture above, Chris snapped me while I was  looking for scorpions on the edge of the Sahara.

By a curious chance, my talk, like a scorpion, has a sting in its tail.

Saturday, 26 November 2011

A stand for microscope slides and cover slips

Here's a quick practical idea.

I wash and dry my microscope slides for re-use, and I needed somewhere to dry the slides. I planned to sit them in saw-cuts in a piece of "2x1" (that's "41x19 DAR" to purists), but the saw I was using did not make wide enough cuts for the slides to fit.

So I used panel pins to hold the slides, and the saw cuts hold the cover slips. The pictures tell the story.

In this shot, that's the washing dish at the back. I blot them on the paper towel then leave them in the rack.


Here is the timber on my work bench, with the saw cuts made.







Just for a neater job, I marked up the alignments for the pins and drilled starter holes.








Then I tapped in the pins.










And finished the job.  That meant cutting off the length that was to be the stand: I usually work with a large piece and trim it when I am ready.





Here it is in use, but because I use circular cover slips, I quickly discovered that when I moved my stand, the cover slips rolled out.

Cover slips are thin, very thin—and they crunch under foot.  This was not a Good Thing, so clearly, Something Had To Be Done!  You can sort-of see what I did here.


But it's clearer in the last shot.

Then  I took some scrap plastic from my scrap box (it was part of a no-longer-needed plastic L plate.)  I cut two strips, and glued them along each side with Superglue. The top of each strip is flush with the top of the stand.

(The other solution would have been to get square cover slips, but I already had these.)


And that was that.

Friday, 25 November 2011

Hunting the elusive tardigrade

My thanks go here to Dr Sandra Claxton who taught me a great deal about tardigrades (actually, she taught me just about everything I know!)


Occasional and skippable one-paragraph commercial announcement: the microscopy material appearing in this blog at the moment comes from out-takes from an upcoming book.   Once you have read this, you will see why we decided to leave this bit out of Australian Backyard Naturalist (which by the way, you don't need to buy or read to follow any of this).

The thing is, this sort of work is advanced stuff, because these animals aren't easy to find: the biggest tardigrades are 1 mm long, the smallest are only 0.4 mm (400 microns). That means you probably won't see them without a hand lens, and you certainly won't see any real detail without a microscope, but tardigrades are everywhere.

Even under x20 with a dissecting microscope, tardigrades are small wriggly blobs, just visible enough to pick up with a brush or a needle, to transfer onto a well slide.

Under a high-power microscope, you will be able to see that they have eight legs, each one usually ending in a claw: only the soil tardigrades are clawless.


Sometimes, the two hindmost legs may be curled up under the body, but after you have seen a few tardigrades, you will learn to recognise the curved claws on the legs. The individual shown here has its two hind legs almost hidden.

The name 'tardigrade' means 'slow walker', but their common name is 'water bear'. Tardigrades are found almost everywhere, from high mountains to deep in the sea, but the easiest ones to catch are the ones that live on or under the bark of trees or among lichens and mosses. You can also find them in leaf litter sometimes.

Some tardigrades drink the juices from plants, but others are hunters, and experts can tell the hunters at a glance, because they have a big pharynx. Tardigrades eat mosses, fungi, protozoa, nematodes, rotifers and even other tardigrades.

Tardigrades are hard to classify, but they seem to be a sister group to the velvet worms and the arthropods. They have no respiratory organs, because they are small enough just to absorb oxygen through their skins. They have a 'straight-through' digestive system, and under the microscope, you can usually see their digestive glands, but not much else.

What you need:
A tree to scrape bark from, a paint scraper, a jar, a coarse sieve, a fine sieve, a wash bottle (see my last post in this blog) some containers and a dish.

What you do:
Scratch some bark fibres off a tree with the side of the blade of a paint scraper, or gather up some moss or leaf litter. You have probably just collected your first tardigrades. Leave this material to soak in water overnight to knock the tardigrades out.

Summary: sieve the damp bark in a coarse sieve. Wash it with a wash bottle and discard the large stuff in the sieve. Take the material that went through the sieve and run it through a cloth sieve to get rid of fine stuff. The tardigrades and similar-sized fragments are on the cloth and can be washed into a jar.

Run the water and bark through an ordinary kitchen sieve, and catch the water in a jug. The tardigrades will now be in the jug, but separated from the big bits of bark.  Note that my "jug" was an ice cream container.



Leave the jug to stand for 30 to 60 minutes, and then strain the water through a 40 micron mesh. If this is hard to get, use a square of an old silk blouse or even a piece of linen. A piece of stocking or pantihose is too coarse, at around 400 microns, the size of a large tardigrade, but even that will catch some, if it is not stretched too tightly.


I have a trick for making filters: I use a wood chisel and a hammer to cut out the flat top of a screw cap, in order to make a sieve funnel: look at the picture above, and the one to the left to get the idea, but I will come back to this trick in a later post, because it can be used for all sorts of things.

Note inserted October 2014: that post has finally been written, and is online.

This second stage separates the tardigrades from the really small stuff in the water.










Then you need to wait patiently as the water goes through, leaving a glug of plant fragments and hopefully a few tardigrades on the cloth.  










The end result after straining is complete.










Then you turn the sieve over and run some water the other way, to wash the tardigrades and anything about their size off the sieve and into a small amount of water. The best way to do this is with a wash bottle.





After that, you just need to search carefully through the remnants in a shallow dish, to see what you can spot moving around. Expect to find all sorts of surprises in there, along with the tardigrades, including large protozoa, nematodes and small mites at the very least. Leave the dish completely still and look for any movement in and under the bits of litter and sand grains. At first, you probably won't see the tiny wriggling shapes without a microscope, but once you know what to look for, a good hand lens will reveal the larger tardigrades.

Another way of catching them

A 'Dust Buster' or other portable vacuum cleaner can save you a lot of work. Fit one with a clean bag and use it to sample tree trunks, lichens and moss mats near waterfalls. You can use it to pick up mites, springtails, beetles, flies, bark lice (book lice) and small spiders. Ian Kinchin, who invented this method, said it was particularly useful on tardigrades.

You need to have a white dish or ice cream container, large enough to let you shake the vacuum cleaner bag into it, banging it with your hand to shake off any small passengers that are hanging on. Then tip the contents of the dish into a holding jar.

In my attempts so far, I think I have used moss that was too dry, so I only got small numbers of mites from the moss. One day, I plan to try two things: first, I want to choose a moss mat and clean it of all twigs, and then 'vacuum' it after rain; second, I want to try watering a moss mat with bottles of water, waiting a few minutes and then 'vacuuming' it. One day, maybe, but you can always beat me to it: if you do, please post a comment here.

Tardigrades are tough! You can find them 6000 metres up mountains and 4000 metres down in the oceans, and on the ground, all the way from the poles to the equator. They can survive being frozen below -200° C for several days, they thrive in boiling hot springs and they can even be heated to 151° C for several minutes. They can also live for a century without water, and for longish periods without oxygen, even in a vacuum, and they can survive huge doses of radiation. People used to say that after a nuclear war, only cockroaches would be alive, but the tardigrades will do even better!

And the moss mat flushing method


Seen at close range, a moss mat offers lots of handy hiding places for small life forms.

In a very real sense (as you will see, once you think about scales), the moss mat is a sort of uniform jungle, though the shot on the right also shows a couple of capsules, the devices from which mosses release spores that can blow around and carry the moss genes to a new place.


In many places, like this bare sandstone slab, to the north of Sydney, a moss mat may be the only cover around. There will be something living there, though probably not much.  Moss growing near a stream or a waterfall or even in a damp alley is a better source.

To catch them, you need a good light source, a Petri dish or a saucer, some moss and some water and/or 70% alcohol. I prefer using just water, so the animals live and I can release them later. The alcohol would kill them.

If you slowly add water to a moss mat in a bowl, some of the animals will climb up into the dry. Adding 70% alcohol with an eye dropper has the same effect, but you need to watch out for fire, and avoid breathing the fumes.* Do this in a flat white dish in the open, pick up the animals with an eye-dropper, and put them in a large amount of fresh sea water to reduce the risk to them—and you!

Remember that tweezers are bad news for animals. You should use a paint brush to pick up any animals you want to mount on a slide for closer examination, and always use well slides to avoid crushing the animals.

If you look carefully, I am told, there may even be tardigrades, but it hasn't happened for me yet.
___________

* By the way, a small amount of methylated spirits in some sea water will flush all sorts of animals out of dried seaweed on a beach.  I will come to that, some other time.

References

(started April 2015)
My thanks to Thomas Boothby (see comments below) who drew my attention to ISTH, the International Society of Tardigrade Hunters. This is an excellent place to go, and you will get real experts there, as opposed to this fumbling enthusiast.

* * * * * * *
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 accepted 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 is covered under the tags Descants and Curiosities, while stuff about small life is under Wee beasties.


Making a wash bottle

I know I said I would do tardigrades next, but for that, you need a wash bottle.  You can buy one (hard to find but expensive) or you can make one—and the design adapts to make a water sampler bottle as well. A wash bottle can squirt upwards to flush material out of upside-down sieves and containers.

The easy way to get one is to take either a drink bottle or a detergent bottle with a 'pop-up' lid and use that, as is. You can use these to squirt upwards or downwards, but it's messy, and if you want a gentle and controllable water flow, you need a proper wash bottle.

You will need a soft plastic bottle (I use a one-litre milk bottle) with a screw-top lid, about 30 cm of 3 mm (internal diameter) plastic tubing, a drill with a bit about the same size as the tubing, some thin iron wire, some gaffer tape (that's duct tape if you are American) and a safe place to use the drill.

The tubing has to reach the bottom of the bottle inside, and come part of the way down on the outside, as you can see from the third picture, so work out the length you need for your bottle. Choose a drill bit that makes a tight hole for your tubing: I use a 13/64" (5 mm) drill bit, but you need to test this for your tubing. Use an old cap and drill several test holes to choose the right size that gives a tight fit.
The wash bottle looks like this when it is assembled.

Then take the cap off the bottle, drill a single hole in it and feed the tubing through the hole until there is enough to reach the bottom of the bottle.

After you have fitted the cap onto the bottle, wind a piece of wire around the tubing so the tubing will take and hold whatever shape you give it. Then cover the sharp ends of the wire with gaffer tape, add water to the bottle, and you are ready to go. You can also use a small amount of tape around the tubing to make a tighter fit where it goes through the cap: this trick also works for pooters.

The wash bottle, ready to use.
(Pooters are in the book, so I won't be describing those here, but I have covered them elsewhere.  Look around and you may find me.  If that seems too hard, try this Youtube clip, posted by my publisher.)

Still with me?  Good.

You can adapt the same design to make a water sampler bottle. I will cover this briefly, because you can experiment with this yourself.

You don't want the water coming out when you squeeze, so if you are holding the bottle right-way-up there should only be a short length of tube inside the bottle. You can also use a long tube attached to a stick to draw up samples from deeper water, but remember to squeeze the bottle before you put the tube in the water, so bubbles don't chase off the wildlife or stir up the sediment too much.

With a bit of thought, you can probably use the same design for a simple one-tube pooter for catching ants.  You need a fairly tight seal where the tube goes through the bottle cap, and I sometimes use epoxy resin glue to seal the tube in place. I advise you to get adult help to experiment with epoxy resin. Work outside, don't breathe the fumes, and try not to get it on your skin. Epoxy resin isn't that dangerous, but play safe!

And now we are ready to tackle the tardigrades.  Next post, I promise!

Tuesday, 22 November 2011

Looking at green slime and hay infusions

A note: the microscopy material that I am putting in this blog at the moment was originally written for, but then deleted from, the upcoming (due out May 1, 2012) National Library of Australia publication Australian Backyard Naturalist, which is directed at readers aged about 10-14.  The majority of them might have trouble accessing or using a microscope, but if they can manage that, well, the information is here to allow them to go the extra mile, so to speak.  You don't need the book to use what you find there.
Click here if you are curious, otherwise, ignore this link.
That's the end of the commercial

The most unlikely puddles can turn out to be home to an amazing range of tiny plants and animals. I like to search puddles beside streams, but I also like searching the scrapings from reeds in a pool or a dam. Things like that often hold surprises. Every pond is a jungle, with animals eating plants and animals eating animals, though some of the smaller animals specialise in collecting scraps and bits. Nothing is wasted in an ecosystem—that's why it's called a system!

Basic microscopy equipment. From the top: backed razor
blade, eye-dropper, dissecting needle, forceps (tweezers),
Pasteur pipettes, brushes.

You will need a microscope, slides (and one well slide), cover slips, a dissecting needle to lay the cover slips down, a medium size camel hair brush, and an eye dropper or a Pasteur pipette (this is an eye dropper with a long drawn-out point). You may also need advice on how to use a microscope, but as soon as you feel ready, start looking at the water fleas or whatever else interests you. Remember Rule 1 of being a naturalist: the most interesting questions are your own questions!



You also need some live material.  Most ponds and still water will develop a collection of life over time, and if you take some green slime, you will normally take a good sample of this life, along with the "slime", otherwise called pond scum, which is actually an assortment of algae and quite a few motile forms that may or may not be animals in the strict sense. The methods for culture outlined in my last entry are probably more useful than going out to collect stuff from dubious drains.

The mistake most beginners make is putting too much material on a slide. Your microscope works when light shines through the stuff on the slide, and your cover slip has to lie down flat. If there is too much gunk, you can't see through it, and the cover slip does not sit down properly. When that happens, you end up with air bubbles which make it even harder to see details.


This is more the sort of thing you want to see. (I cheated here: this is another part of the same slide, but that doesn't matter, so long as there are less clumpy bits.  Practice makes perfect!

To master this trick, get a tiny amount of green slime from a pond, put it in a drop of water on a slide and add a cover slip. When you look at your first slide, it will always be too dark to see anything. Make a second slide, and this time, use even less slime, and spread it out with two needles or two pairs of forceps.

Even then, some parts of the slime will be too thick, so move the slide until you are looking through a less populated part, and focus on it under low power. Then, if something looks interesting, put it in the middle of the field of view, and move to a higher power. (This instruction will make more sense once you are used to working with a microscope.)

Take small amounts of "slime", and spread it out with a brush. If you have a fish tank, some of the most interesting gunk comes from the filter. Stir this up a bit, and then take some of the material with an eye dropper and put it in a Petri dish. With a black background and a strong light, you should be able to see if there are any tiny animals in the water. If there are, use a Pasteur pipette to take a sample. If there are no visible animals, use a flat slide, if you can see moving animals in the gunk with your naked eye, use a well slide.

When you switch to high power, you may see Paramecium (you can see one of these in the video which is in my last entry), some nematodes, or some of the other larger animals that live in green water. Paramecium is a single-celled animal with the common name "slipper animal", because they are shaped a bit like a slipper. They are ciliates, which means they move along by beating large numbers of short hairs called cilia.

The large ciliates feed on bacteria, yeasts and algae, all of which have smaller cells. Keep an eye out also for filamentous algae like Spirogyra. That one is easy to spot under the microscope because it has spiral chloroplasts.

(This one is not Spirogyra, but all filamentous algae are formed when a series of single cells link up to form a thread like this. This one is x400.)


Paramecium cells range in size from 50 µm to 350 µm, which means the largest ones will be just visible with a hand lens, if you are lucky. I see them most often as large dark blurs that shoot across my microscope's field of view, out of focus, when I am looking at something smaller, like diatoms.

Diatoms are single cells most of the time, but some of them form filaments, ribbons and even colonies. They have cell walls made of silica, and diatomaceous earth is formed from the remains of long-dead diatoms. There are probably 100,000 different diatom species around the world.

There are three diatoms in this picture, but two of them were on different levels and so more blurred. Viewed at x400.


To identify your algae, you will need a dichotomous key to the green algae, or the filamentous green algae. To search on the web, it will probably be better to enter a search string using the technical name for the green algae: <key freshwater chlorophyta> or <chlorophyta identification freshwater>. Leave out the "angle brackets".

Filamentous algae are called 'moss' and 'pond scum' when they attack fish ponds and tanks, where they often go out of control and end up dying to leave a decaying mass, riddled with bacteria which use up all the oxygen in the water, killing the fish. They are actually long strings of algal cells. Some of the most interesting filamentous algae can be found where water runs through channels in a rock. The cells in the filaments are too small to see with a hand lens, but you can see detail even under low power with a microscope.

Most of what you see will be algae. I suggest that you look up the following on the web and become familiar with the appearance of Spirogyra, Volvox, Scenedesmus, Nostoc and Chlamydomonas. As you track those down, you will probably see other familiar algae as well. One thing you are sure to find is rotifers.  I will finish this entry with a bit about them.

Rotifers

Oops! As oldsalt19 has pointed out, this is a protozoan
called Vorticella. Colour me absent-minded!

Rotifers are also called 'wheel animals', a name that refers to their cilia, which beat continuously as a way of catching food, and look like spinning wheels. You could call them worms if you wanted, but 'worm' can mean almost anything, because it isn't a scientific term.

Under the microscope, rotifers are always moving, trying to catch food. You can also see one in motion in the video in the previous blog entry.

Rotifers are quite small, about 100 to 500 microns (0.1 to 0.5 mm) long, though a few are as large as 2 mm. Mostly, they are found in fresh water, and the ones you will probably see are the sessile forms, the kind that grip onto something and stay there, but other rotifers are free-swimming, and some others 'inchworm' their way around.

From their size, you might think that they are single-celled protozoa, but they have multiple cells, an alimentary canal with a pharynx (think of it as a mouth) and an anus. They mainly eat single-celled algae like Chlorella, Euglena and Chlamydomonas.

Rotifers have a very simple nervous system. In many species, males are rare. No males are known at all for any member of the family which includes the genus Rotifer, and they reproduce by producing eggs which have a full set of chromosomes. (This is seriously specialised: look up <bdelloid rotifer diploid egg> for more information.)

To get some rotifers to study, collect some pond water and stand it on a window-sill in moderate light for a few days. The rotifers will collect near the top, where there is more oxygen, so you can pick them up with a pipette or an eye-dropper. You will also find some attached to filamentous algae and other bits and pieces in 'green slime'.

They also live in moss mats. If you soak a piece of moss mat in water and then squeeze it out over a bowl, you will generally find nematodes and rotifers. The method is supposed to produce tardigrades. I have never found tardigrades so far, but lots of rotifers!

I will come to the tardigrades next time.  They are cute, quite hard to catch, but here is the proof that I managed to catch one. This was so ferociously difficult that it had to go, because it would just put too many younger readers off. Still, for those with grit, determination, or just a plain stubborn streak, I will tell my tale and share what I learned.

* * * * * * *
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 accepted 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 like this, all about small life is under Wee beasties.

Monday, 21 November 2011

Culturing algae, green slime and animalculi

videoThis 6-meg video was taken through my microscope.  It's there mainly to show you how some of the things you see actually look under the microscope. The cup-on-a-stalk low on the screen is a rotifer, the long whizzing thing is a Paramecium, and the little round things are little round things.

They all grew in one of my algal culture systems, so there is every chance that you could see the same things with your cultures as well.

In this entry, I will show and tell you how to set up cultures, and explain some of the tricks I use to get good results. There is no best method, but there are lots of good methods.

My suggestion is that you take my ideas and work them up into what you want.

First, here is a method that I used to use, some 40 tears ago. I had a series of large glass bottles, all with air coming from a small aquarium pump. At the end, the air bubbles through a tray.

I would put each inlet tube (the bubbling ones) just below the water surface, to avoid too much strain on your air pump. You need clear bottles so light can get in to make the algae grow. Looking through towards sunlight on a sunny day, you may be able to see either a Paramecium, or an Amoeba in your cultures, so look these up on the web.  You may also see some very tiny crustaceans: look for tiny moving dots.

Because I often wanted to open one of the bottles, I used plastic tubing for the connections, even back then, because glass tubing would have been too dangerous. In the end, though, I decided that this method was unnecessary.

Here is the kit that I had set up, just outside my study window, in full sunlight. The old film canisters were used to collect water from puddles and streams, so there were always new organisms likely to be getting into the system. At this time, I needed mosquito wrigglers, and I used the flat dish to collect some.

The main extra was that I added some fertiliser to the water samples.  Not a lot, just enough to make sure the plant life had something to work with.  Note that some of the containers are back in the shade: the dish, in particular, got up to 60°C on some summer days, and that was another variable.

A hay infusion
The traditional method, though, is to prepare a hay infusion.  This is the traditional name given to this sort of thing, dating back to the days of people's horsepower having four legs. You don't really need hay: any plant matter that has been lying around will do.


You will need an old saucepan, a bucket, a few PET bottles to store cultures in.  PET bottles are those clear plastic bottles that soft drinks come in.  Use the 1.25 litre size, but never fill past the point where the sides slope in near the top, because you want a large water surface for gas exchange.

(One handy trick: if there are small crustaceans in a bottle, if you fill the bottle almost to the top, oxygen levels will drop, and they will all cluster in the top 5 mm (where there is more oxygen), where you can catch them with an eye dropper.)

Use the saucepan to boil some grass or chopped-up leaves in water.  After it has cooled, let the water stand in a bucket for a day or two. Use the water to cover some cut grass or leaves, and put the container in a warm place, out of direct sunlight. You can speed things up if you add a few small slops of water from a pond or a slow stream, or some of the cloudy water from the bottom of a vase of dead flowers.

I am usually in a hurry when I need algae, so I have a few extra PET bottles, half full of water and assorted gunk samples that serve as sources. I can use these to get a new culture started, or I can draw samples.

A few small water samples from random pools and ponds can help boost the variety in your infusion containers. One of the best ways to collect water samples is to get a used washing-up liquid container, the sort that has a pop-up/pop-down lid. Wash this out thoroughly and take it with you when you go walking.

When you find some nice water, pop the lid open, squeeze some air from the bottle, turn it upside down, push the top under water and unsqueeze to get a sample. You can increase your animal catch by taking water from close to plants, rocks or the bottom. Always take a plastic bag to wrap the wet bottle in after you have taken a sample. I now always wash my hands after taking samples like this, using bottled 'hand-wash', and I recommend that you do the same.

After that, you will need a bit of patience, because the best results and the biggest surprises come from cultures that have been going for a while.  In the next entry, I will talk about some of the stuff you can see.

Here's one example, at a magnification of x400. It's what we call a filamentous alga, but which one it is, I'm sure I don't know.  Like you, I'm just a hobbyist, dabbling where I feel like dabbling.

Making wet mounts

A note for those coming in part-way through:

The material that is going into my writing blog (that's this one) is currently being drawn from edited draft material that I wrote for Australian Backyard Naturalist.  This book (ISBN 9780642277428) will be published by the National Library of Australia in May 2012, and because we were shaping the concept as we went, I wrote rather more than twice as much material as we were able to fit into what will, in fact, still be quite a large book.

I wanted to add microscopy, but it would be somewhat beyond the reach of part of the target audience, and it tended to involve sharp blades, worrisome stains and a fair amount of expense.

I still believe in the idea of introducing young people to microscopy, so I have taken about 11,000 words of draft, and I am slowly feeding it in here.  Remember that you really need to read the entries in reverse to get the sequences and sometimes the logic.

Right now, we are up to wet mounts: putting something on a slide and covering it with a cover slip.

Now read on . . .

* * * * * * * * *

It is possible just to look at a drop of water on a slide, but you can see more through a flat surface, and that means using a cover slip of very thin glass to flatten the water out. You will almost certainly break a few cover slips and cut yourself at least once. When you are starting out, wear safety goggles to protect your eyes, and practise very hard at being gentle with the cover slips.

By now, I hope you will have tried making the odd thin section, cut using a home-made microtome. If you want to skip past that and look at tiny life forms, I will come to some ideas in my next entry. For now, think this sections, placed on a microscope slide, spread flat with a brush, a needle or forceps, but needing a cover slip to be added, without trapping any air bubbles.

Here's how you do it, first as a simulation:


Making wet mounts: a simulation using a knitting needle and a microscopy plate, about 10 cm across. The change of scale meant the operator had to hold the "cover slip" to stop it slipping. Notice how the needle always touches the "slide" and comes out gradually.


And now, the real thing.  There is nothing in the water here, but there could have been a slice of carrot, a piece of onion skin, some blood or skin (if you had been really careless with the razor blade)—the options are endless.

You need to avoid air bubbles, and that means putting the cover slip down so it touches the drop of water on one side, while holding the slip up with a dissecting needle on the other side. Then you slowly pull the needle out, keeping the needle down at an angle of maybe 20°, so that the cover slip comes slowly down on the drop, and the air underneath is pushed sideways.

The surface tension effects stop the cover slip from skipping away as the cover slip is lowered: notice how the water front pushes across. I used too much water in this one, so as to make it show out. If I planned to look at this mount, my next step would be to sop up some of the water, using the corner of a face tissue.

You now have the basics: the next post will be rather longer, and will concentrate on what you can find in "green slime". There's a whole new world waiting for you out there!

Thin sections and a microtome

One way to beat the depth-of-field problem is to cut thin sections of things so all of the parts of your specimen are at the same level, just one cell thick. You can do this using the sort of razor-blade that has a strong backing edge. These are very sharp and dangerous, so get adult advice before you start. Thin sections can be mounted in water under a cover slip, and they will let you see cells, though without stains (we'll come to those later) you won't see much internal detail.

The basic microtome. I inserted a plug of carrot into the wing
nut, which is fitted back to front onto the bolt. You need a
dish of water to put the sections in, and something to pick them up.
Before long, you will realise that these 'thin sections' are usually wedge-shaped, and you can see better detail at the thin side of the wedge. If you want a thin section that is even all over, you need a microtome. Microtomes are expensive, but there is a way to make one almost for zilch—well, mine cost just $2! All I needed was a razor blade and a matching half-inch Whitworth wing nut and bolt (some hardware is still sold in "old" units).


ADULT SUPERVISION is essential for this one—risks include cuts and possibly broken blades hitting the eye. You will need a bolt with a wing nut to match, safety goggles, a safe very sharp blade and an old cutting board.

The equipment you will need. Note the paintbrush. the dish, the
chopping board. the $2 microtome and the backed razor blade in
particular. You will need the needle later, to mount a section.
For your first attempt, cram a short piece of carrot (or celery) into the wing nut's threaded hole, then slip the bolt into the wing nut back-to-front, with the 'wings' at the end nearest the bolt.

When you go to slice a section of carrot, you will see why the nut has to be this way around. Once the bolt has a grip on the nut, put on the safety goggles, get the cutting board and use the blade to trim off all the carrot that is sticking out of the nut.



As the bolt slowly moves into the nut, the carrot in the threaded hole is slowly pushed out on the other side, and if you slide a sharp blade across the flat surface of the wing nut, you will cut that tiny bit off, producing a thin section that can be mounted on a slide. Trim off any bits that are sticking out of the nut.

Now you can start. Turn the nut slightly, so a tiny amount is pushed out of the threaded hole in the nut, and slice downwards.


Hold the apparatus as shown on the left: this shot was posed to let you see a section coming off. Don't use it as a guide!








This is the way to hold the equipment.

Notice in two of these shots the Petri dish with water in it, that you drop the sections into as you cut them. A saucer or even a jar lid will do just as well.









 You lift the section with the brush . . .
 Carry it across to the dish . . .

And put it in the water. While you are learning, put the slices in a dish of water repeat the operation until you master the method and throw away your first attempts. Examine the sections in a wet mount if you wish, but now you are ready to section difficult stuff like leaves.

For small items like leaves or stems, you will need some scrap polystyrene foam to wrap around what you are sectioning. You can also use cork or a piece of carrot or potato for this, anything that grips tightly on the leaf or stem.

If you read up on professional microtomes, you may see references to using wax instead of foam, but I recommend against this. It is hard to make a water mount of a waxy section, because wax and water don't mix. Most professional reference books recommend using very toxic chemicals to dissolve the wax, so polystyrene foam is safer than wax. Do some experimenting first: and remember that a piece of foam larger than the hole can always be squeezed and 'screwed' into the nut, once it is wrapped around the leaf or other object.


Put the wing nut on the bolt again, with about one full turn of the nut on the bolt, then fill the empty portion of the nut with whatever you want to section. If you really want to section a leaf in wax, prop the bolt upright in an old jar or can, poke the leaf in, and then drip candle wax in, until the leaf is surrounded with wax and leave it to set.


The sections will usually be wedge-shaped, but if you choose to look at the thinnest part, you will be able to see something like this shot, which is one of the sections you see being cut above. It isn't perfect, but it's a good start.









Of course, I have left out a big step: actually getting the section on the slide and putting a cover slip on top without any bubbles. That is called Making a wet mount, and I will deal with that next.

For comparison, here is a small portion of the first-ever drawing of cells, done from a section of cork, cut by Robert Hooke with a pen-knife in 1665 (this link gives you the whole thing in Wikipedia).

Saturday, 19 November 2011

A bit more about microscopes and hand lenses

Sherlock Holmes and hand lens,
as seen in the Strand Magazine.

I have now ferreted out all of the discarded text on microscopy that might have gone in Australian Backyard Naturalist and didn't. The rationale was simple: we had about three times as much text as would fit in a standard book, and something had to go.

Realistically, not everybody has a microscope, and many of the readers might not have access to a microscope. Then there was the consideration that microscopy is not a core element of the publishers' mission statement.

That said, it is part of mine, and in any case, you can see quite a lot with a hand lens, as Sherlock Holmes knew.

Mind you, the Holmesian lens wasn't up to all that much, delivering a magnification of around x10, but there is a whole variety of hand lenses that we can call into service, and the one on the lower right can deliver about x20, as can a (fairly expensive) achromatic hand lens.

Let's remember, though, the $50 USB microscope that I mentioned in my last entry. That sort of thing ought to be in most people's reach, and the sorts of things I will be writing about here are all reachable with one of those: and some of them can be done, less satisfactorily, with a hand lens.

To be honest, some of the shots I am using here have come from one of the microscopes you can see on the right, and when it comes to thin sections, you can see the difference.  The gadget lying between them is the semi-professional USB camera that I use with those: its slips into the top of the microscope, in place of the objective.

I stopped half-way through the last paragraph to have dinner, and part of the meal was raw onion ( which we happen to like). Anyhow, I filched a small portion of onion and peeled the epidermis off it so I could make a slide and run it through the$50 toy.

From left to right, we have two versions on the $50 microscope, then a x100 and a x400 from the monocular microscope using the good light on the expensive camera.  The main difference, I think, is in the light source.  Still, anything you buy today will be better than the lighting Robert Hooke had to put up with in the 1660s.

We've come quite a long way since then, and perhaps the most important is that young Robert had to do his own drawings (or some of them: tradition has it that Sir Christopher Wren did some of the work).  We just use the computer, easy-peasy!

In the next entry, I will look at a neat method of cutting this sections using a nut, a bolt and a razor blade, and how to mount a slide with a cover slip.