3.5: Motility Agar - Biology

Because the flagellar staining procedure often produces poor results in the hands of novices, other tests for motility (and the presence of flagella) have been developed. One type of test involves using a semi-solid medium that allows motile bacteria to penetrate.


  1. Obtain a deep of motility agar.
  2. Using an inoculation needle, stab straight into the deep about 2/3 of the way down and out the same pathway as quickly as possible with your assigned bacterium.
  3. Incubate the tube for at least 48 hours.
  4. After the incubation period, examine your tube.


If the organism is motile, growth will ‘fan’ out from the stab or the entire tube may look cloudy. If the organism is non-motile, growth remains confined to the stab line.

Negative Motility

Tests for Bacterial Motility: Procedure, Results

The motility test is used to determine whether an organism is motile or non-motile. Motile organisms contain flagella which helps them to travel beyond the point of inoculation. Motile bacteria are generally bacilli although a few motile cocci do exist. Motile bacteria move with structures called flagella (a few exceptional bacteria move with the help of axial filaments, which cannot be seen in the microscope). Motility test helps us to differentiate between genera and species of bacteria.

The triple sugar- iron agar test employing Triple Sugar Iron Agar is designed to differentiate among organisms based on the differences in carbohydrate fermentation patterns and hydrogen sulfide production. Carbohydrate fermentation is indicated by the production of gas and a change in the colour of the pH indicator from red to yellow.

To facilitate the observation of carbohydrate utilization patterns, TSI Agar contains three fermentative sugars, lactose and sucrose in 1% concentrations and glucose in 0.1% concentration. Due to the building of acid during fermentation, the pH falls. The acid base indicator Phenol red is incorporated for detecting carbohydrate fermentation that is indicated by the change in color of the carbohydrate medium from orange red to yellow in the presence of acids. In case of oxidative decarboxylation of peptone, alkaline products are built and the pH rises. This is indicated by the change in colour of the medium from orange red to deep red. Sodium thiosulfate and ferrous ammonium sulfate present in the medium detects the production of hydrogen sulfide and is indicated by the black color in the butt of the tube.

To facilitate the detection of organisms that only ferment glucose, the glucose concentration is one-tenth the concentration of lactose or sucrose. The meagre amount of acid production in the slant of the tube during glucose fermentation oxidizes rapidly, causing the medium to remain orange red or revert to an alkaline pH. In contrast, the acid reaction (yellow) is maintained in the butt of the tube since it is under lower oxygen tension.

After depletion of the limited glucose, organisms able to do so will begin to utilize the lactose or sucrose. To enhance the alkaline condition of the slant, free exchange of air must be permitted by closing the tube cap loosely.

Enzymatic digest of casein (5 g), enzymatic digest of animal tissue (5 g), yeast enriched peptone (10 g), dextrose (1 g), lactose (10 g) sucrose (10 g), ferric ammonium citrate (0.2 g), NaCl (5 g), sodium thiosulfate (0.3 g), phenol red (0.025 g), agar (13.5 g), per 1000 mL, pH 7.3.

There are many variations of the catalase test. The methods discussed here are the slide method, tube method, and tube (slant) method. Other methods such as the semiquantitative catalase test for the identification of Mycobacterium tuberculosis, the heat-stable catalase test for differentiating Mycobacterium species, the capillary tube method, and the cover slip method are not elaborated here. (5)

  • Using a sterile inoculating loop or sterile wooden stick, get a small amount of an 18- to 24-hour old microbial colony, and place it on the clean, dry glass slide.

Avoid picking up any agar, especially if using agar that contains red blood cells. Red blood cells contain catalase, and would yield false-positive results.

  • Using a dropper, add a drop of 3% hydrogen peroxide to the bacterial colony on the microscope slide.
  • Observe immediately for the formation of bubbles. Placing the slide against a dark background facilitates easier observation. (5)

Picture 2: Slide method for catalase test.
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Individual interphase microtubules in control Dictyostelium cells are exceptionally motile, displaying a variety of rapid movements that appear to be driven by both plus end– and minus end–directed motors (Figure 1 Supplementary Movie 1 Kimble et al., 2000, Koonce and Khodjakov, 2002). Such movements include simple bending and straightening, flailing back and forth in a whip-like manner, and arcing along the cell cortex. Forces acting on the microtubules appear able to originate at any point along the entire microtubule length. Overexpression of the dynein motor domain introduces an additional motile component: the entire microtubule array appears to circulate through the cytoplasm and along the inner surface of the cell cortex, at rates up to 2.5 μm/s (Koonce et al., 1999 see also Supplementary Movies 3a and 5c). We used a laser microbeam to sever the connection between the centrosome and the trailing comet tail of microtubules in these 380K cells (Figure 2 Supplementary Movies 2a–d). In 15 of 18 cells, centrosomes immediately cease their movement, suggesting that much of the motility is driven by force exerted on the trailing microtubules. The microtubule arrays in the three cells whose centrosomes continued moving were less focused, and we could not be certain that most of the microtubules had actually been severed. Control irradiations in the cytoplasm, adjacent to or in front of the centrosome had no effect on this movement. In some instances after cutting (Figure 2D), microtubules were reorganized on the other side of the centrosome and began to drive motion in the direction opposite the preirradiation movement.

Figure 1. Interphase microtubule movements in a control cell. An arrowhead marks the plus end of a single GFP-labeled microtubule in Dictyostelium. In the first four frames (0–30 s), the plus end follows a whip-like bending trajectory through the cytoplasm, in a direction consistent with the application of kinesin-like forces to the microtubule. In the last four frames (33–45 s), the same microtubule end now leads, in a direction consistent with the application of dynein force. The microtubule itself maintains a constant length (13.3 μm) throughout the sequence. The position of the centrosome is marked with an asterisk in the first panel (t = 0). Time indicates seconds of observation scale bar, 2.5 μm.

In nearly every case, the laser-mediated liberation of minus microtubule ends resulted in the disassembly of the trailing arrays. However, in a few cells (e.g., Figure 3), the microtubule array appeared tightly bundled and did not immediately disassemble. In these cells, we found that the microtubule bundles themselves were capable of motion, independent of the centrosome. This result indicates that the centrosome itself is not required instead, the force responsible for the motility acts along the microtubule bundle (Supplementary Movie 3b).

To distinguish whether the force is due to molecular motors or microtubule-end dynamics, we measured assembly/disassembly rates for Dictyostelium microtubules. The goal was to determine whether this organism's tubulins have any unusual kinetic properties that could contribute both to the lateral movements in control cells and to the centrosome movements in the 380K cells. We restricted our analysis to those microtubules whose entire length remained in focus throughout the observation period. Our records revealed that while transitions between growing and shrinking are extremely rare, Dictyostelium microtubules do undergo dynamic instability (Figure 4A). Using the laser microbeam, we also created free plus and minus ends of single microtubules. Except for those in tightly bundled microtubule arrangements (as described above), free minus ends of microtubules in Dictyostelium are unstable (Figure 4B). In a number of cases, we could readily follow this depolymerization and thus generate rate measurements for minus end depolymerization (Figure 4C). We saw no evidence for tubulin assembly at free minus ends of microtubules. Plus ends were more stable: only half of the newly created plus ends depolymerized to varying degrees, whereas the other half remained stable throughout the observation period (Figure 4D).

Figure 2. Laser-mediated cutting of motile microtubule comet tails. (A–D). Four examples of 380K cells in which the trailing GFP-labeled microtubule bundle is separated from the leading centrosome. The first panel in the A–C sequence shows a “before” view the second panel shows the cell immediately after (or during as in C) laser cutting of the microtubule tail. The two rows in D show an extended sequence of a binucleate cell, containing two microtubule arrays. The arrowhead marks the site of laser ablation. In all cases shown here, the centrosome stopped movement upon severing of the microtubule bundle. Note in D that the second microtubule array movement is unaffected by the ablation and that in the last few panels (153 s and beyond), the unaffected microtubules on the side opposite to the cut have begun to organize and push the centrosome toward the bottom of the frame. Time is displayed in seconds of observation scale bar, 5 μm.

Our measurements of microtubule end dynamics (assembly and disassembly) are shown in Table 1 these averages are comparable to kinetics seen in GFP-labeled mammalian microtubules (also included in Table 1 Rusan et al., 2001). In contrast, the rapid movements of individual microtubules in Dictyostelium occur at rates averaging 1.1 μm/s, approximately four times faster than could be explained solely on the basis of assembly or disassembly. There were no rate differences between what we interpreted as plus end– or minus end–directed force production, a finding consistent with other measurements of organelle trafficking in Dictyostelium (Table 1). These results strongly imply that the rapid microtubule and centrosome movements in control and 380K cells are largely due to molecular motor activities, rather than to unusual kinetics of assembly.

Table 1. Average rates and comparisons of microtubule assembly, disassembly, and motor-driven movements in control cells

What is Semi Solid Media?

Several techniques are used to observe and detect the motility of bacteria. Among them hanging drop method is one such method. However, it has several disadvantages such as the tedious nature of the method, uncertainty of the results, difficulty of identifying the motility when only a few cells are motile, need of active or fresh cultures etc. Hence, scientists have developed semi solid media for above purpose. Semi solid media are microbial culture media that are prepared to add less amount of agar (solidifying agent at 0.2 to 0.5 %) to observe motility of bacteria. Semi solid medium was first introduced by Hiss in 1982 for the purpose of distinguishing typhoid and colon bacilli.

Figure 02: Stab Tube

The results of the semi solid media are macroscopic. When motile bacteria are inoculated to stab cultures that were prepared using Semi Solid Media, a diffuse zone of growth along the inoculation line of the stab can be clearly observed. It eliminates the overlooking of motility if only a few are motile.

Everything you need to know about agar

Agar (or Agar Agar), sometimes referred to as kanten, is a gelling agent coming from a South East Asian seaweed. It is used for scientific purposes (in biology for instance), as a filler in paper sizing fabric and as a clarifying agent in brewing. Agar can also be used as a laxative (it’s 80-percent fiber) and as an appetite suppressant.

And it’s of course an amazing culinary ingredient. It’s a vegetarian gelatin substitute, a thickener for soups, in fruits preserves, ice cream and others desserts.

Where can you find agar?

Agar is available in health food stores, in supermarket that carry health food lines, in Asian grocery stores and online.

Health Benefits

Agar has no calories, no carbs, no sugar, not fat and is loaded with fiber. It’s free from starch, soy, corn, gluten, yeast, wheat, milk, egg and preservatives.

It absorbs glucose in the stomach, passes through digestive system quickly and inhibits the body from retaining and storing excess fat. Its water absorbing properties also aids in waste elimination. Agar absorbs bile, and by doing so, causes the body to dissolve more cholesterol.

A great substitute to gelatin

Agar is the perfect substitute to traditional gelatin. It’s made from a plant source rather than from an animal one. That makes it suitable for vegetarian and vegan diets, and other diet restrictions.

Agar has no taste, no odor and no color, which makes it pretty convenient to use. It sets more firmly than gelatin, and stays firm even when the temperature heats up.

Though agar is a great substitute to gelatin, don’t expect the same results when replacing gelatin with agar in a recipe. First, it doesn’t give the same texture. Gelatin can give a «creamy» texture whereas agar gives a firmer texture. And agar is much more powerful than gelatin : 1 teaspoon agar powder is equivalent to 8 teaspoon gelatin powder.

How to use Agar

- The most important thing to know is that agar needs to be first dissolved in water (or another liquid like milk, fruit juices, tea, stock. ) and then brought to a boil. It will set as the ingredients cool down. You can not add agar flakes or powder as it is in your food.

- You should definitively follow the package directions and the recipe to determine which quantity to use. But here is a basic rule you can adapt : use 1 tablespoon agar flakes to thicken 1 cup of liquid, and 1 teaspoon agar powder to thicken 1 cup of liquid.

Here is the basic «recipe» to use if you can’t boil your liquid directly.


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#4 Hanging drop method

In a wet mount, there is a possibility that the motility of bacteria is hampered. Hence, a hanging drop method is more reliable. Such a method enables you to clearly observe the motility of bacteria including the size and shape.

What is the principle of hanging drop method?

A tiny drop of bacteria is hung from the middle of the cover slip into the cavity slide. It is then observed under the microscope using the oil-immersion objective. The movement in the medium and its surrounding indicates that the bacteria are motile.

On the other hand, if the bacteria remain calm in the medium, it means that the bacteria is non-motile. (1, 4, and 6)

  • Cavity slide
  • Coverslip
  • Lubricant (petroleum jelly)
  • Broth culture of bacteria
  • Immersion oil
  • Loop
  • Microscope
  1. Petroleum jelly is applied around the clean and dry cavity slide.
  2. The loop is put over the flame and allow to cool. Using the sterilized loop, a bacteria is taken from the broth culture.
  3. A drop of suspension is put at the coverslip’s center.
  4. The slide is inverted and put on the coverslip. The two are pressed gently to seal the cavity. Make sure that not a single part touches the drop.
  5. The slide is inverted so that the drop hangs into the cavity.
  6. The slide is then clipped to the stage and will be examined under the low power objective of the microscope. (6, 8, and 9)

With the hanging drop method, the bacteria will be checked for its motility, shape, arrangement, and size.

Watch the video: low motility (January 2022).