This weekend we have had guests – and they asked me what tap water looked like under the microscope. We filled 18 Eppendorf 1.5ml centrifuge tubes with tap water and centrifuged them at 10000 revs/minute for 10 mins. We then pipetted off the top 95% of the water and moved the remaining water into a single tube and centrifuged it again at similar settings. Removing the top 90% of that tube and then putting the bottom few drops onto a slide and using x40 objective revealed that there is was a small amount of grit and organic debris only. We could not see bacteria. This suggests that our tap water is pretty clean and safe.
Cellular Turbulence. Rhys and I went with the family to the Big Bang Show at the NEC in Birmingham. On the Zeiss stand were a number of microscopes – and on one of them some Elodea showing chloroplast movement around the cell. This pond weed has particularly mobile chloroplasts and the site is amazing. This movement is referred to as cyclosis or cytoplasmic streaming.
See the photo and video below – wow! I wonder what the small things are, much smaller than chloroplasts? Organelles or parasitic protozoa or bacteria? Magnifications here using x32 and x63 objectives so bacteria would show up at this magnification.
Some facts about chloroplasts:
Chloroplasts are organelles, specialized compartments, in plant and algal cells. The main role of chloroplasts is to conduct photosynthesis, where the photosynthetic pigment chlorophyll captures the energy from sunlight and converts it and stores it in the energy-storage molecules ATP and NADPH while freeing oxygen from water. They then use the ATP and NADPH to make organic molecules from carbon dioxide in a process known as the Calvin cycle. Chloroplasts carry out a number of other functions, including fatty acid synthesis, much amino acid synthesis, and the immune response in plants. The number of chloroplasts per cell varies from one, in unicellular algae, up to 100 in plants like Arabidopsis and wheat (from https://en.wikipedia.org/wiki/Chloroplast)
The average chloroplast is about 3 µm (micrometers) in diameter. In one square millimeter of the surface of a leaf, there are about half a million chloroplasts (from www.answers.com/Q/What_is_the_size_of_a_chloroplast)
Chloroplasts in vascular plants range from being football to lens shaped and as shown in Figure 1, have a characteristic diameter of ≈4-6 microns (BNID 104982, 107012), with a mean volume of ≈20 μm3 (for corn seedling, BNID 106536). In algae they can also be cup-shaped, tubular or even form elaborate networks, paralleling the morphological diversity found in mitochondria. Though chloroplasts are many times larger than most bacteria, in their composition they can be much more homogenous, as required by their functional role which centers on carbon fixation. The interior of a chloroplast is made up of stacks of membranes, in some ways analogous to the membranes seen in the rod cells found in the visual systems of mammals. The many membranes that make up a chloroplast are fully packed with the apparatus of light capture, photosystems and related complexes. The rest of the organelle is packed almost fully with one dominant protein species, namely, Rubisco, the protein serving to fix CO2 in the carbon fixation cycle. The catalysis of this carbon-fixation reaction is relatively slow thus necessitating such high protein abundances (from http://book.bionumbers.org/how-large-are-chloroplasts/)
Components of cells seen in photos and video:
Today’s photos and video:
If you want the wow factor go straight to the videos at bottom of page using x63 objective! I am quite excited by the views with the x63 objective below as this is first time I have used it so successfully with this microscope. The slide is turned upside down and Kohler illumination has been achieved using my “new” (second hand off ebay) NA 0.9 bright field condenser with Zeiss 475638 illuminator collimation tube (at least I think that is what it is for!).
The slide was prepared using free hand sections of Elodea leaf using razor blade put on slide with drop water and covered with cover slip. Edges of cover slip help firmly to slide by electrical insulating tape strips and then slide turned upside down and put on microscope stage (upside down as Zeiss IM microscope is an inverted microscope).
Photo of Elodea leaf section (cut free hand with razor blade) x32 objective, bright field, Zeiss IM microscope, showing cell walls and chloroplasts:
Photos of Elodea x63 objective bright field, now also shows small inclusions much smaller than chloroplasts – in later videos these are shown to move as well as chloroplasts:
Videos – first two videos are with x32 objective, bright field:
Videos – next videos used x63 objective, bright field:
My Zeiss IM microscope came set up for phase contrast. Although phase contrast is an amazing technique, the condenser was limited in its ability to be used to set up Kohler illumination.
Kohler illumination is a method of illumination of microscopic objects in which the image of the light source is focused on the substage condenser diaphragm and the diaphragm of the light source is focused in the same plane with the object to be observed; maximizes both the brightness and uniformity of the illuminated field (http://medical-dictionary.thefreedictionary.com/Kohler+illumination)
To achieve Kohler illumination with the Zeiss IM or IM35 microscopes, I needed to obtain one of Zeiss’ bright light condensers. The manual showed the microscope in use with a flip top condenser so I purchased one of these from ebay together with the extension tube and condenser diaphragm shown in the manual – had to wait a bit until one was available.
Success! I can now focus the diaphragm edge in the same plane as the image of the slide, improving illumination and contrast to the maximum available for the microscope…..at least in theory – and seems to work today when I tried it.
Carl Zeiss 0.9 NA Swing Flip Top Condenser (below). With this arrangement, I am able to open the diaphragm up to the full field of view:
Condenser mounted on microscope – both with flip top lens in and out (below):
Image of condenser diaphragm edge shown against the background slide – I have closed down the diaphragm somewhat to show the edge (below):
I also tried mounting one of my other Zeiss condensers. This one has bright field, phase and dark field options. It achieved focus although the condenser had to be much closer to the slider, but, even when fully open, the diaphragm could not illuminate the whole field evenly (below):
The following photos show a piece of plant stem I cut up from a pot in our kitchen this evening. I don’t know what the plant was. It shows may chloroplasts and cell walls.
x20 bright field:
x20 objective Phase Contrast I annulus:
x32 objective Phase Contrast I annulus:
The following two photos are both taken using the x32 objective and phase contrast I annulus. The long thin features look like bacteria but they did not move so I wonder if they come from the plant? I wonder if they might be raphides. Raphides are oxalate of calcium needles secreted by some cells. Here, the edges of my blade may have cut an epithelial cell full of raphides throwing them on the cut surface, in a similar way to that experienced by Walter Dioni in his post on http://microscopy-uk.org.uk/mag/indexmag.html?http://microscopy-uk.org.uk/mag/artfeb04/wdstem.html
This sample was collected from our garden 7 days ago and kept in an open jar of water.
The contents of the jar had separated into a top layer of moss floating on the top, an intermediate layer of very cloudy water and a bottom layer of debris on the bottom of the jar. I have tried to sample all three layers in the pictures below.
x20 objective bright field sample from bottom of jar – debris layer. This shows large numbers of bacteria.
x32 objective bright field bottom debris layer:
Moss 7 day culture bottom jar layer video x32 objective Phase I annulus:
x20 phase contrast I debris layer bottom jar:
x32 objective phase contrast I debris layer jar:
x20 objective phase contrast I cloudy liquid layer between debris on bottom and floating moss – I am not convinced that this is phase contrast even though I labelled it as such – looks like bright field to me now:
x20 bright field liquid layer between debris and moss – video:
x20 bright field one single moss plant from the floating moss on top of the jar. If you look carefully you can see hundreds of bacteria surrounding this plant:
Tardigrades are water-dwelling, eight-legged, segmented micro-animals. They were first discovered by the German zoologist Johann August Ephraim Goeze in 1773. The name Tardigrada was given three years later by the Italian biologist Lazzaro Spallanzani. They have been found everywhere: from mountain tops to the deep sea and mud volcanoes (Wikipedia).
Tardigrades, often called water bears or moss piglets, are near-microscopic animals with long, plump bodies and scrunched-up heads. They have eight legs, and hands with four to eight claws on each. While strangely cute, these tiny animals are almost indestructible and can even survive in outer space. Tardigrade is a phylum, a high-level scientific category of animal. (Humans belong in the Chordate phylum — animals with spinal cords.) There are over 1,000 known species within Tardigrade. Water bears can live just about anywhere. They prefer to live in sediment at the bottom of a lake, on moist pieces of moss or other wet environments. They can survive a wide range of temperatures and situations (https://www.livescience.com/57985-tardigrade-facts.html)
I went looking for tardigrades today in St Michael’s church graveyard in Lichfield, Staffordshire, UK. No success – sadly – so you won’t see tardigrades in the photo and video below. However, the samples I obtained from moss on gravestones, some lichen off trees and a sample from a wood chipping pile, revealed a range of life shown in the video below.
Vorticella on a commercial stained slide, viewed using Zeiss IM microscope at different magnifications. Damian and I have previously seen Vorticella live in local pond water samples – see previous posts.
I used the Bresser MikOkular camera in “new” trinocular head (second hand from ebay) – this differs from previous trinocular head in that this is the one that is recommended in the Zeiss IM microscope handbook. I noted that the previous head, although it works, has small black ring around outside of field of view that I assume means field stop is too small for scope. This new one does not have this. This new one also provides 23mm ocular attachment on trinocular port, into which the Diagnostic Instruments adapter fits directly without needing a clamp.
I also tried out a dark field condenser on microscope today – did not work well – not sure why – so photos below are back to the phase condenser that came with the microscope, used without phase annulus (i.e. in bright field mode).
x63 objective (slide upside down so light only has to go through coverslip in this inverted microscope – 63x objective has only limited working distance). This is a panorama of 17 panes, joined using Microsoft’s Image Composite Editor:
This is where the epi-illumination technique comes into its own. The following pictures are of a UK 1£ coin – showing up surface relief differences and tiny scratches that are not otherwise visible to naked eye.
I have used the white balance adjust function in the Bresser MikroCamLabII software (camera control software) to remove the effect of the colour tinge imparted by the mirror in the filter cube on the microscope.
Following photos x4 objective:
Helicon Focus stack – interestingly this does not appear to have improved a great deal on above best focus image: