MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
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Unit II: Bacterial Cellular Structures and Morphology
Activities:
2.3 Motility Test (Ex 5)
2.4 Wet Mounts & Observing Live Bacteria (Demo)
3.1 Smear Preparation & Staining (Ex 6)
3.2 Gram Stain (Ex 7)
3.3 Endospore Stain (Ex 8)
4.1 Introduction to Brightfield Microscopy (Demo)
Appendix 2.1: Notebook Pre-Lab Preparation Example
p1
p2
p5
p7
p9
p 12
p 18
2.3 Motility Test:
The TTC Motility agar is a semi soft agar used to determine if a bacterium is motile or not. The agar
contains the chemical 2,3,5-triphenyl tetrazolium chloride (TTC) which turns into red formazan dye
due to presence of respiring bacteria. The media is inoculated by carefully stabbing a loopful of the
bacteria into this agar. Motile bacteria should be able to swim away from the stab line thus an overall
red-pinkish hue will be observed in the whole tube after one or two days. Non-motile bacteria will
stay in the stab line and a red color will be observed only along the stab line. Having made
microscopic observation of different live bacteria, you should be able to predict your TTC motility test
results.
EXERCISE 5:
Materials:
1. Inoculating needle
2. Bunsen Burner
3. 24 hour tubes of Proteus mirabilis, Bacillus megaterium, and Staphylococcus epidermidis
4. 4x TTC motility agar tubes
Procedure:
1. For each of the three broth cultures of bacteria, label a TTC Motility stab tube.
2. Using sterile technique, dip the tip of the inoculating needle into the broth culture.
3. Stab the needle in the TTC Motility stab agar to the bottom of the tube. Stab straight in and then
back out along the same path of entry. You want a clean puncture, not a gouge.
4. Incubate the tubes for 24 to 48 h at 37ºC.
5. You should examine these tubes over the next two days; taking notes on your results and
conclusions.
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
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6. How do your results from the microscopic observation of live bacteria compare to the TTC Motility
stab results? Don’t forget to discuss any conflicting results. You may wish to draw some opinion on
which assessment of motility is most reliable.
2.3 Wet Mounts & Observing Live Bacteria:
One of the most elementary skills used in identification of a bacterium is microscopic observation.
Microscopic observations stained and unstained bacteria can reveal a variety of characteristics used for
identification. These characteristics include bacterial cell size and shape, cellular arrangement,
motility, Gram reaction, and the presence of a capsule, endospore, or inclusion bodies.
We will start the microscopic examination by preparing a wet mount (next lab period you will
learn different staining techniques). Wet mounting is the preparation of unstained live bacteria. The
wet mount will allow us to observe cell size, shape and arrangement, the presence of endospores of
inclusion bodies, and determine if the bacterium is motile.
Size: Length and width, or diameter, are the size measurements we’re concerned with. To perform
these measurements one needs to have a calibrated micrometer installed in the ocular. The
micrometer allows you to estimate size dimensions.
Shape: Basic shapes include spherical (cocci, coccus), rod (bacilli, bacillus), spirals (spirilla,
spirillum), curved rods (vibrio), flexible multi-curved spirals (spirocheates) and elongated filaments
(filamentous). Some bacteria are pleomorphic (unusual varied shapes) or prosthecate (stalks or
appendage-like structure emanating from the main body of the cell).
Cell arrangements (Figure 2.1): Cells may be found as singles, pairs (diplo; diplococcus,
diplobacillus), strings (strept; streptococcus, streptobacillus), cubic shaped packets (sarcina),
irregular clusters (staph; staphylococcus, staphylobacillus), or aggregations forming long cords.
Motility: You must remember that in a wet mount there are turbulent currents generated in the liquid
as you focusing and moving the slide. This means cells may be seen streaming, or sweeping, in one
direction across your field of view. Furthermore, once any turbulence calms, Brownian motion will
produce an effect in which the bacteria looks “shimmering” or vibrating in the slide. These two types
of observations are not to be mistaken for motility. True motility is evident as bacteria moves in a
different direction from the normal flow of liquid on the slide. In other words, it should look like York
at rush hour … traffic in all directions at once!
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
Figure 2.1. Bacterial cell arrangements (Fig. 3-12; Leboffe and Pierce, 1999).
DEMONSTRATION
Microscopic Observation of Live Bacteria:
Materials:
1. Phase-Contrast Microscope (only one; we’ll have to share)
2. Microscope slides and cover slips
3. Inoculating loop
4. Bunsen Burner
5. 48 hour plates of Proteus mirabilis, Bacillus megaterium, and Staphylococcus epidermidis
6. sterile water
7. 24 hour tubes of Proteus mirabilis, Bacillus megaterium, and Staphylococcus epidermidis
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MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
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Procedure:
1. Set up the microscope and make sure you have all material you need at hand. Also, be prepared to
draw and write down your observations in your lab book.
2. Prepare a wet mount using the bacteria from the broth provided. To make a wet mount you need to
add two or three loopfuls of a broth culture onto a clear slide and then carefully place a cover slip on to
the slide (it will be demonstrated by instructor). Alternatively, if your source of bacteria is a plate
place two or three loopfuls of water on a slide then pick a small amount of bacterial growth from the
plate and mix it well with the water. Make sure you dissolve all clumps and cover it with a cover slip.
Make one mount at a time; otherwise, the preparation may dry out before you have adequate
time to perform observations.
3. When first viewing a slide, begin with the low power objective (may start with 4x and move to 10x)
and try to focus on something within the slide that you can see with the naked eye such as the edge of
the cover slip or better yet an air bubble on the slide. Once you have found something with the 10x
objective close the aperture diaphragm slightly to increase the contrast. Now, switch to the 40x phase
contrast objective (remember to flip the dial on the condenser to the annular stop for phase contrast).
You will probably have to adjust the aperture diaphragm again. The 40x is the most important
objective you will use for examining live material. Spend some time observing things using this
objective. It is usually much easier to observe living cells using the 40x phase contrast objective than,
40x simple brightfield or the 100x oil immersion
4. Take note of shapes, sizes, arrangements, and look for the presence of motility for each of the
three different bacteria and your own pet bacteria.
5. Now proceed to 100x magnification by moving the 40x objective out of the way, take the
microscope out of phase contrast, and place a single drop of oil directly on the cover slip, place the
100x objective in place making sure that the oil is making full contact with the lens. . The cover slip is
relatively thick and can move when you bring the 100x objective into place so be careful. Adjust focus
with the fine focus drive and readjust the aperture diaphragm to give maximum contrast. Now move
around and examine the live material at 100x magnification. Record your observations in your lab
notebook.
6. Dispose of the wet mounts as directed by your instructor. Clean the immersion oil from the
100x objective. Place the low power objective in place, wrap the cord around the base and take the
microscope back to its proper location in the cupboard.
Summarize the observations you made. Compare the strains observed. You will use these data
in the future in comparing other techniques, such motility agar testing and staining procedures.
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Unit II: Bacterial Cellular Structures and Morphology
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3.1 Smear Preparation & Staining:
In the previous laboratory you were able to visualize unstained live bacteria using phase
contrast microscopy. Today we will be using different staining techniques to visualize bacteria and
some of their structural features using bright field microscopy.
Why do we need to stain bacteria? One reason is to increase the contrast between the
bacterium and its surroundings. Unfortunately, many of the staining techniques result in the death of
the bacteria and may also lead to some change in structural features. These changes may include
shrinkage and changes in shape. What other feature cannot be evaluated in dead bacteria?
Before staining the bacteria, a culture preparation must me smeared on a glass slide and air
dried. For most staining procedures, once the preparation is air dried the cells are “fixed”, glued or
bound, to the glass by application of heat from your Bunsen burner flame. Heat fixing the preparation
binds the specimen to the glass slide so that it will not be washed away during the staining steps. The
staining itself consists of the addition of chromophore (a colored molecule or compound) which is
usually dissolved in some liquid (could be simply water). With few exceptions, most of the stains used
are cationic (positively charged, or basic). This means that the chromophore will combine strongly
with the negatively charged cell surface components (Figure 3.1). Usually the area of the smear is
flooded with 2-3 drops of the stain which is allow to stay for a couple of minutes, and then the excess
stain is washed away with water, finally the slide is blotted dried and examined under the 100x oil
immersion
objective.
Figure 3.1. Simple stain mechanism, whereby charge-charge interactions bind basic chromophores to
the net negative charges of the bacterium (Fig. 3-1; Leboffe and Pierce, 1999).
Some types of Staining:
1) Simple Stain:
This is usually a very fast and simple procedure where the fixed preparation is stained for a couple of
minutes and then rinsed and blotted dry. Some of the stains commonly used in simple stain are:
methylene blue, crystal violet, safranin; all of these stains are cationic, i.e. basic.
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
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2) Negative Stain:
The chromophore of a negative stain is negatively charged (anionic or acidic) hence it will not
combine with the same charge cellular components. In fact, these acidic stains are repelled from the
cells surface. Because the acidic stain will bind to the glass slide but not the cells, the result is a dark
stained background and clear, colorless, cells. This staining technique is very useful because it can be
done without heat fixing thus the distortion of the bacterial cell is minimized. Examples of
chromophores used in negative staining are: eosin, India ink, nigrosin, and fuchsin.
3) Fluorescent Staining:
Fluorescent stains are used to enhance observation sensitivity over that for colored stains. The
technique requires a fluorescent microscope, which exposes the specimen to ultraviolet (UV) light.
Here the specimen is stained with a fluorescent dye that will fluoresce, i.e. emit a visible light color,
upon excitation with the UV light. One common stain used is acridine orange which stains nucleic
acids. You will be able to use fluorescent microscopy later on in the semester.
4) Differential and Structural Staining:
These are staining technique designed to differentiate between types of cells. One of the most
important is the Gram stain which differentiates cells on the basis of their cell wall structure. Other
stains include the acid-fast stain, which is important in the identification of the bacterial genus
Mycobacterium. Among commonly used structural stains are the capsule stain, flagella stain, and
Schaeffer-Fulton spore stain.
Viewing Stained Slides:
Observe the stained slides initially at 10x magnification so that you can find the material and get it into
focus. Then switch to the 40x objective, the material should still be into focus with a slight adjustment
of the fine focus knob. Finally, move the 40x objective out of the way and place a single drop of oil
directly onto the stained area of the slide, then move the 100x objective back into place.
NEVER MAKE AN INTERPRETATION OF A STAINED SLIDE UNTIL AFTER YOU HAVE
EXAMINED IT AT 100X MAGNIFICATION USING IMMERSION OIL!
EXERCISE 6:
Preparation of Heat-Fixed Smears:
Simple stain, Gram, and endospore staining techniques will require a heat-fixed smear of bacteria
while capsule stain will not.
1) Make sure you start with a very clean glass slide. You can free a slide from grease and dirt by
using a paper towel to rub 2-3 drops of ethanol onto the slide and allow it to dry well.
2) Use a wax pencil to faintly mark the bottom of the slide with a 1.5 x 3 cm ellipse.
3) Place a small loopful of water onto the marked circular areas.
4) Using aseptic technique, touch a colony with your loop to collect some biomass (not too much!).
5) Place the loop with the bacteria onto the drop of water and mix the water and the bacteria. As you
mix, smear the suspension out in a thin layer. The thinner film of cell suspension the better.
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Unit II: Bacterial Cellular Structures and Morphology
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6) Let the smear air dry. This may take several minutes, so you may want to prepare all the smears
for one staining technique at once (e.g. all the smears of bacteria to be Gram stained can be
prepared at once). Remember to keep track of what is on what slide. Besides labeling the slide I
recommend placing them on a paper with their label written beside each to help keep track. Also,
you may use a small piece of masking take on the top side with a number that you will write in
your lab notebook to identify what organisms is in what slide.
7) To HEAT FIX take a completely dry smear and pass it through the flame with the preparation side
(bacterial smear) facing away from the flame (up side). Usually, three slow passes will heat the
slide enough. You should be able to touch the slide to the back of your bare hand (without glove).
The slide should feel warm but not uncomfortably hot. If the slide feels too hot the bacteria has
been fried and you need to make a new smear.
3.2 Gram Stain:
Method overview:
The Gram stain technique separates bacteria into two main groups. Gram positive are the bacteria
that will retain the color of the first (primary) stain used (crystal violet). Gram negative bacteria are
those that assume the color of the second stain (safranin).
The difference in staining occurs because bacteria differ in the structural composition of their cell
walls. Gram positive bacteria have thick peptidoglycan layers and tend to retain the crystal violet
even after a decolorizing step involving a ethanol rinse. They stain purple, and stay purple even when
the red safranin stain is added later. The addition of an iodine wash, called a mordant, after crystal
violet staining helps to complex the dye with Gram positive cell wall compounds Gram negative
bacteria have thin peptidoglycan layers surrounded by an outer membrane. Their cell wall will initially
get stained by crystal violet, but then is easily decolorized with ethanol in the decolorizing step, which
results in removal of any purple. The cell wall is then stained (counterstained) red when the safranin is
added. The exact reasons why the two types of cell wall react so differently to the Gram staining
technique is not well understood but it appears that the presence of different components on both walls
and the thickness of the peptidoglycan layer play important roles.
The decolorization is the differentiation step in Gram staining. Again, this step is basically a 95%
ethanol rinse. In Gram negative bacteria, decolorizing dissolves the lipid bilayer of the outer
membrane and allows for rapid removal of the iodine crystal violet complexes. In contrast for the
Gram positive bacteria, decolorizing dehydrates the thick cell wall, which helps retain iodine-crystal
violet complexes. Of course to long a rinsing with ethanol will eventually rinse away the crystal
violet even from Gram positive cell.
In summary, there are four critical steps, each followed by a water rinse (Figure 3.2):
1) Crystal Violet (purple)
2) Iodine Mordant
3) Ethanol Decolorizing
4) Safranin (red)
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
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EXERCISE 7
Materials:
24 hour old plates of Proteus mirabilis, Bacillus megaterium., Staphylococcus epidermidis.
Gram stain reagent kit
Wax pencil, slides,
staining trays.
Steps 1&2
purple
purple
Step 3
purple
Step 4
red
purple
Figure 3.2. Gram stain procedure, showing the result at each step for both a Gram positive and a
Gram negative cell. Note the first two steps, crystal violet and iodine mordant; they have been
combined into one for this diagram. (modified Fig. 4-1; Leboffe and Pierce, 1999).
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Unit II: Bacterial Cellular Structures and Morphology
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Procedure:
1) Prepare a fixed smear of the bacteria to be stained as explained in Exercise 8 above.
2) Flood the slide with crystal violet for one minute to completely cover the smear(s). Note that you
do not have to cover the entire slide just the area where the smear is.
3) Wash with water. Remove the excess water by gently taping the edge of the slide on a paper
towel. Don’t squeeze the stream of water directly on the smear area; rather, target above the smear
and let the water gently rinse over the smear.
4) Flood the slide with Gram’s iodine (the mordant) for one minute.
5) Wash with water and remove the excess as directed earlier.
6) Decolorize with 95% alcohol. Flood for approximately 10-30 seconds, or for a thin smear 10-15
or 20 seconds. CAUTION: Excess flooding of ethanol may eventually decolorize Gram positive
cells as well.
7) Wash with water as indicated in step 3
8) Flood the slide with safranin for one minute
9) Water rinse as in step 3
10) Drain and blot dry with the special blot paper (bibulous paper) provided.
NEVER RUB THE SLIDE!
11) Wait until the slide is completely dry before trying to view them under the microscope.
12) Record your observations of shape and on your lab notebook; include sketches.
13) Summarize your results on Gram reaction, cell shape and cell arrangement, and discuss how the
later two observations compared to those made on live cells of the same bacteria in Exercise 6.
3.3 Endospore Stain:
Bacterial endospores are small oval structures that are produced as a dormant or resting stage by some
bacteria of the genera Bacillus and Clostridium. The bacterial endospores are resistant to desiccation
(drying), heat, radiation, and disinfectants. Thus, they provide a means of preservation for bacteria
facing an unfavorable environment. Unfortunately, this property has been capitalized on by
bioterriorists who have successfully mailed endospores of Bacillus anthracis to spread forms of the
infectious disease, anthrax.
The size and the location of the spore within the vegetative bacterial cell are used to help
differentiate different species of bacteria. The spores are either larger or smaller than the vegetative
cell. They are located within the cell either terminally (at the end of the cell), subterminally (near the
end of the cell), or centrally (in the middle of the cell).
Schaeffer-Fulton endospore staining is a differential staining technique that allows the
visualization of released spore, cells with spores contained within, and vegetative cells with no spores.
Any spore appears green and any vegetative component or cell stains red. Because spores have a heat
resistant spore coat the primary stain (Malachite Green) must be driven into the spore by the use of
excessive heat. In addition, the heat will help to disrupt the wall of any vegetative cells which prevents
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them from retaining the stain during the rinse process. The vegetative cells are then counterstained
with safranin red (Figure 3.3)
EXERCISE 8
Materials:
Malachite Green Stain (5% in water)
Safranin counterstain (0.5% in water)
24, and 96 hour old cultures of Bacillus megaterium
Hot plate, 50 ml Beaker with water and boiling chips.
Procedure:
1) Prepare a fixed smear of the bacteria to be stained as explained in exercise 7 above. In order to get
the best results, take biomass from the edge of colony growth on the 24 h plate and from the center
of a colony on the 96 h plate. HINT: which has ample nutrients for vegetative growth? Which is
starving?
2) Flood the slide with Malachite Green, and place on the beaker of boiling water using the slide
holder so you don’t scald your hand.
3) Heat the flooded slide so that the stain begins to steam. When steam begins to rise from the
stain, add more stain to replace the evaporated stain. Thus, you will have to keep replacing the
stain to keep the slide flooded. You want the stain to steam for 5 minutes …DO NOT LET THE
SLIDE DRY OUT …
4) After the 5 minutes of steaming drain the slide and rinse with water for 30 seconds. Remove the
excess water by gently taping the edge of the slide on a paper towel.
5) Flood the slide with safranin and allow it to stain for one minute.
6) Drain the slide and rinse thoroughly in water. Remove the excess water as indicated in step 4.
7) Carefully blot the slide and allow it to air-dry.
8) Examine the slides and determine the relative size and the location of the spore. Note which age
culture was most prolific in spore production. Write down your observation in your lab notebook,
including sketches.
9) Discuss the observations in the context of growth and nutrient supply (availability).
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
Figure 3.3. The Schaeffer-Fulton spore stain method (Fig. 15.1; Benson, 1994).
4.1 Introduction to Brightfield Microscopy:
Some terminology you should become familiar with:
brightfield microscopy
total magnification
ocular lens
objective lens
numerical aperture
resolving power
oil-immersion len
parfocal
stage
condenser
Some of the exercises in this and other subsequent microbiology courses require that you are
competent in the use of microscopes. The microscopes that we use in this course are brightfield
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compound microscopes. This same basic compound microscope can have additional optical
accessories added to permit other interchangeable applications, such as darkfield and phase-contrast
microscopy. An overview of phase-contrast microscopy is included below; however we will reserve
its discussion until the next lab unit when we’ll put is to use.
Nomenclature and Parts: (Figure 4.1)
Let’s look at the anatomy of the binocular (two eyepieces) Brightfield compound microscope
and discuss the function of each part, beginning from the eyepieces and progressing along the light
path to the light source, or lamp.
Figure 4.1. Brightfield compound microscope parts (Fig 2.3; Prescott et al., 2002).
The total magnification of any compound microscope is the product of the ocular (eyepiece)
lens and the objective lens. Ocular lens magnification is typically 8x, 10x, 12.5x, or 15x. Depending
on your individual eyesight, you may have to adjust the ocular lens focus on usually just one eyepiece.
Also, depending on the size and shape of your head you may need to make an interpupillary
adjustment. It is probably a good idea to note these adjustments for the specific microscope you plan
to use in all labs. Students in other courses will use the microscope during the semester, so your
microscope oculars may require adjustment every period.
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Unit II: Bacterial Cellular Structures and Morphology
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The revolving piece above the specimen stage has several objective lenses attached. The
objective magnification is marked on each objective, 4x, 10x, 40x, or 100x. The numerical aperture
(also called field flatness) of each objective is the value found after the slash (/) e.g. 40/0.25 means it is
the 40x objective and its numerical aperture is 0.25. The larger the numerical aperture, the greater the
resolving power, i.e. the ability to distinguish between two closely packed objects. The other set of
numbers (e.g. 160\ 0.17) refers first to the length of the mechanical tube (160 mm) used with these
lenses and then the correct thickness (0.17 mm) of the cover slip to be used. If a dash “-“ is present
instead of the later number, it means that the objective can be used to view specimens without a cover
slip. The word PHACO written on the lens means that it can be used for phase contrast microscopy.
In addition to the values mentioned above, each objective has a “working distance” which is the
distance between the specimen and the front surface of the lens when it is in sharp focus.
Let’s look at the each objective. The 4x objective is called the scanning objective it has a working
distance of 17-20 mm. It is wise to start with this objective when a new slide is placed on the stage.
The 10x objective is called the low power objective and it has a working distance of 4-8 mm. The 40x
objective is also called the high dry and is used most often for wet mounts and hanging drops. It has a
working distance of about 0.5-0.7 mm. The 100x objective is the oil immersion lens and it is marked
with a black circle around the neck. Its working distance is 0.1 mm. As its name indicates, the oil
immersion objective is used with special oil, which has almost the same refractive index as the glass
slide. This objective is usually used for the examination of stained bacteria. Microscopes equipped
with proper objectives will be parfocal, i.e. the specimen stays focused when objectives are changed.
This property allows use to easily find objects starting at low magnification and then “zoom-in”
sequentially with higher magnification and only minor fine focus adjustment.
The stage is the area where you place the slide containing the material to be seen. Just under
the left side of the stage there are two knobs that are use to maneuver the slide laterally and/or
longitudinally. When performing oil immersion, keep the stage free of oil and make sure any excess
oil is wiped from the stage. An oily stage will case your slide to be “sticky” and retard smooth
movement of the slide as you turn stage knobs.
The condenser unit is located under the stage. The condenser contains a lens, called a
“condenser lens”, which gathers the light coming from the lamp so that a cone of light is projected
through the slide and into the objective lens. The condenser can be adjusted in our microscopes to
maximize the numerical aperture of the lens by widening the cone of light; thereby, it increases
resolving power. Some condensers require being on high position for the 100X and down for 40X.
You should not have to ever adjust the condenser, unless you or someone else accidentally moves it.
Please remember that condenser height is not used to control focus, nor it is used to adjust the intensity
of the light. To adjust light intensity, there is a knob on the base of the microscope (below the coarse
and fine adjustment knobs).
The illumination tube is located at the base of the microscope. It contains a lamp and the field
diaphragm (iris). Although adjustment of the field diaphragm is necessary for some of the work
(Kohler illumination), it is usually left totally open.
Phase-Contrast Microscope: (Figure 4.2)
We have a microscope equipped for phase-contrast microscopy, and we’ll use this in
demonstrations in a latter lab unit. The optical accessories added to permit a brightfield compound
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
microscope to perform phase-contrast microscopy are a annular stop and a phase plate which is
present in the objectives used for phase contrast (in our case the ones that have the word PHACO
written on it).
Figure 4.2. Phase-contrast microscopy optics (Fig 2.9; Prescott et al., 2002).
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The idea behind phase contrast is to increase the contrast (light to dark) between the cells and
its surrounding background so that they can be seen live, i.e. without staining. This is achieved by
using two elements: 1) the condenser annular stop, which is simply an opaque disk with a transparent
ring to produce a hollow-cone of light; and 2) the objective phase plate (an special optical disk)
containing a phase ring. Light that hits the phase ring is considered an undeviated light ray, i.e these
rays came directly from the light source and did not hit specimen objects. The phase ring has a special
coating that decreases the intensity of the light by about 75% (and advances its wavelength ¼). The
deviated (deffracted) rays are those that did hit specimen objects. The different density of the
specimen causes rays to be retarded by ¼ wavelength . These, retarded rays pass the phase plate away
from the phase ring, and their property is unchanged. The final effect is that the deviated and
undeviated light rays are about ½ wavelength out of phase with each other resulting on negative
interference, i.e. they cancel each other and result in darkness. A pronounced contrast between the
specimen objects (dark) and their surroundings (light) results. That is to say, the background appears
bright and the object appears dark and well defined.
Operation of the Microscope
BEFORE YOU START! These microscopes are rather expensive pieces of equipment. CARRY THE
MICROSCOPE WITH TWO HANDS, AND HOLD IT CLOSE TO YOUR BODY FOR ITS
SAFETY. Once the microscope is place on the bench, completely unwind the cord from the
microscope and plug it in.
Microscope Set Up:
Familiarize yourself with the microscope. Identify the parts of the microscope:
a) The on/off light switch on the back of the base.
a) The light source and the knob for the light intensity regulation.
b) The lever for adjusting the aperture diaphragm (iris)
c) The stage, the slide holder (object mount) and the drive knobs of the object guide for moving the
specimen
d) The objective lenses (4x, 10x, 40x and 100x) on the rotating objective nosepiece. Rotate the
nosepiece and note that each lens “click-locks” into position.
e) The ocular lenses. Note how the spread of the eyepieces can be adjusted for your own individual
comfort
f) Coarse and fine focus drives.
g) Condenser.
Focusing the Image:
a) Slide a specimen containing microscope slide into the specimen stage.
b) You may start by using the scanning objective (4x) or you may turn in the low magnification (10x)
objective. Notice: the microscope should always be stored with the lowest magnification
objective into position.
c) Switch on the illumination and regulate its intensity by turning knob.
d) Make sure the condenser is on the upper stop near the specimen.
e) Make sure the aperture diaphragm is open.
f) Adjust the distance between the oculars (interpupillary distance) until the image for both eyes
completely cover each other and appear as a single circular image.
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g) Carefully focus on your specimen. * REMEMBER TO WATCH FROM THE SIDE (i.e. NEVER
MOVE THE STAGE UP WHILE LOOKING THROUGH THE OCULARS) as you lower the
objective lens, and focus UP only with the coarse control. If a slide smashes against the objective,
it may cause extensive damage to the lens. Also remember that this microscopes are parfocal
which means that the image should remain in focus when the objectives are changed.
HINT: try to focus on the edge of the cover slip or a small air bubble first. Line it up so the
edge is directly under the objective lenses, and then use the drive knobs of the object guide to
move it slightly as you focus. You may also need to adjust the intensity of the light.
h) Accommodate the distance. Do not get too close when looking through the ocular lenses (your
eyeballs should not be making prints on the lenses). How do you know if you are too close?
When you bring your eyes too close to the eyepiece lenses you may see that the entire field of
view appears to float away into darkness. Another problem that you may find is gunk on the
ocular. To make sure what you are seeing is indeed on the slide; rotate the eyepiece lens a little.
If the dubious object moves too, clean the lens with lens paper. You may also improve the
contrast by moving the aperture diaphragm. This diaphragm needs to be mostly closed before you
will see unstained bacteria under low power.
Using the immersion oil objective requires the used of special immersion oil. A drop of immersion oil
is applied to the slide (either to the surface of the cover slip or to the smear). Swing the 100x objective
into position. The lens should be in full contact with the oil (with no air bubbles). Look into the
eyepieces and adjust focus using the fine adjustment knob only. It is very important that you do not
get oil into the 40x objective lens. If oil accidentally get onto the 40x lens inform your instructor
immediately. Proceed to clean the lens as shown by your instructor.
After the investigation. all surfaces where the immersion oil has been applied must be carefully
cleaned. Use the special lens paper supplied moistened with alcohol or Windex . Place the lowest
magnification objective into position. Wrap up the cord around the microscope and take it back to its
place on the cupboard.
TROUBLE SHOOTING:
if you have problems finding your specimen check these:
1- Microscope is plugged in and light turned on.
2- Objective lens is not clicked into position
3- Slide is not laying flat
4- Double coverslip
5- Slide is place on the stage upside down
6- Iris diaphragm is closed
7- Eye pieces are dirty or unbalanced
8- Light source is set too low or high
9- Objective lens is too far from smear
10- Condenser is too low or high
11- If you can see your specimen with the 10X, but when you turn to 40X you notice that the objective
will touch the glass YOUR SLIDE IS UPSIDE DOWN. This is a very common occurrence. It can be
avoided by clearly marking with a wax crayon the bottom side of the slide where the specimen is
placed (i.e. specimen is in the unmarked side).
MICRB 202: Introductory Microbiology Lab
Unit II: Bacterial Cellular Structures and Morphology
17
APPENDIX 2.1: UNIT II: Samples Lab outlines and flowcharts.
Exercise 8: Preparing Fixed Smears
Obtain two clean glass slides. One of the slides will be use to place a few drops of water

For each slide that will be used for specimen do the following:

1- Use a wax crayon to draw a circle at the bottom part of slide

2- Place a piece of masking tape at the end of the topside of the slide. It will be used to
write the number that identifies the specimen

3- Use loop to transfer a drop of water from one the slide that has water to slide for
specimen

4- Sterilize inoculating loop and allow it to cool

5- Use sterile loop to touch a colony of specimen to be studied

6- Place loop with bacteria sample on the drop of water on slide and mix well. Spread
the mixture well over the slide to allow for faster drying.

7- Set the slide aside to air dry the smear

8- Once the smear is dry fix it by passing the slide (smear side up) quickly through the
flame. Touch the slide to the bare skin of the back of your hand to make sure it is not
too hot. Repeat this twice.
Smears need to be made for the following bacteria for use in staining exercises #9 & 10.
1x
2x
1x
1x
Proteus mirabilis for Ex 9
Bacillus megaterium (24h) for Ex 9 and Ex 10
Bacillus megaterium (96h) for Ex 10
Staphylococcus epidermidis for Ex 9.