Week 8

For week 8, we performed several tests to determine the enzymatic properties of our bacteria. These tests included Methyl Red test (mixed fermentation), Citrate utilization test, Indole (tryptophan degradation) test, Oxidase test, Nitrate Reduction Test, and the Urea hydrolysis test.
To begin, we inoculated test tubes containing the medium of each specific test, and then incubated them until the following lab to determine the results.

The following lab period, upon retrieving our test tubes, we determined that our bacteria had a negative test with Urea, Oxidase, and Citrate, but a positive test with Nitrate, Indole, and the methyl red fermentation test.

 
For the Nitrate test, we took our inoculated nitrate broth tube, and added 5 drops of nitrate reagent A (sulfanilic acid) and 5 drops of nitrate reagent B (dimethyl-alpha-naphthylamine). Our nitrate broth immediately took on a deep red color, indicating a positive test.

For our Indole test, we took our inoculated tryptone broth tube and added 10 drops of Kovac's reagent. A red layer appeared at the tope of the tube, indicating that the test was positive.

Next, we took our methyl red broth tube and poured out half into another sterile test tube for a second fermentation test (Voges-Proskauer Test) We then added 5 drops of methyl red (pH indicator) to the first tube.

Our tube took on a reddish color, indicating a positive test.

We then took the second test tube of methyl-red broth and added 15 drops of Baritt's reagent A (alpha-naphthol) and 5 drops of Barritt's reagent B (KOH) to the tube. Our broth did not change colors, indicating a negative test.
Finally, we performed the oxidase test. Taking our inoculated broth tube for the test, we took a sample using a sterile swab. We then added the oxidase reagent to the swab. There was no change in color, indicating a negative test for oxidase.

In conclusion, through these tests, we concluded that our bacteria:
a) is able to ferment glucose via mixed-acid fermentation, but not through butanediol fermentation (methyl red)
b) does not use citrate as its sole source of carbon and energy (Citrate test)
c) has the ability to split the amino acid tryptophan into indole and pyretic acid (Indole test)
d) does not have cytochrome oxidase (oxidase test)
e) is able to reduce nitrate ions to either nitrite or to nitrogen gas (nitrate test)
f) and is unable to hydrolyze urea

Week 7

Week 7

10/10/13

This week we observed the results from all of the tests we prepared last week.

We determined that our Starch hydrolysis test was negative, thus our bacteria does not have the enzymes necessary to digest starch.

Our casein hydrolysis test (skim milk plate) also was negative.

Our gelatin test was negative 

 

 Our fat (triglyceride) hydrolysis test was negative (our bacterial sample is on the bottom)

 

And finally, our Litmus milk test results:

The white curd at the bottom detects peptonization with an alkaline reaction. Thus, our bacteria has proteolytic enzymes that digest the casein (curd) into peptides and amino acids.

We were then instructed to inoculate test tubes filled with four different mediums to test if our bacteria had enzymes to digest carbohydrates (these test tubes were filled with lactose, sucrose, glucose, and Triple Sugar Iron (TSI) mediums).

Week 6

Week 6
10/3/13

Last Tuesday we inoculated a tube of nutrient broth with our unknown bacteria sample that we received in class. We then incubated the broth solution at 37' C for the remainder of the week. 

We spent today's class streaking the same unknown bacteria over various mediums, to see which enzymes our bacteria expressed. Bacteria can be classified is by these enzymes, bringing us closer to identifying our sample.

We inoculated bacteria in litmus milk

and gelatin,


and streaked it over skim milk, starch, and spirit blue (lipid) agar plates.

We will incubate these samples at 37'C until our next class.

Week 5

Week 5 

9/24/13 - 9/26/13 

in week 5 of class we performed an acid-fast stain, an endospore stain, and determined if our bacteria was motile or not. We first started with the acid-fast staining. An acid-fast stain is used to distinguish the two groups of bacteria based on the lipid content and their cell walls: acid-fast and non-acid-fast. We first made a bacterial smear on a glass slide and then fixed it over a Bunsen burner.  We then placed the slide over a beaker of boiling water. We took a piece of bibulous paper and set it on the slide before dropping carbolfuchsin over the length of the slide. We continued to drop carbolfuchsin on the slide for 3 minutes.

We then removed the bibulous paper and retrieved the slide using forceps. We rinsed the slide using  water to remove excess stain and then a decolorizing solution, adding drop by drop until the color stopped running. We then immediately rinsed the slide to remove the decolorizing agent. We then covered the slide with methylene blue for 2 minutes.

Then we rinsed the slide with water to remove the excess coloring. After blotting the slide with pieces of bibulous paper we examined the smear under the microscope

upon examination we determined that are bacteria was a non acid-fast due to the methylene blue stain

We then were given a tube with motility test medium. Using aseptic technique, we took a sample of our bacterial culture with an inoculating needle and then stabbed it into the test medium. We then placed it in the incubator at 37' C until the next lab period.

The following lab, we worked on preparing an endospore stain. This stain is used to determine if a particular bacteria has endospores. We again fixed a smear of bacteria on a glass slide. We then placed the smear over a beaker of boiling water. Taking a piece of bibulous paper and placing it over the smear, we dropped malachite green over the smear. We let the stain set for 6 minutes while continuing to add stain as it evaporated.

 We then discarded the paper and retrieved the slide using forceps. We rinsed the slide with water to remove the excess dye. We then covered the smear with safranin for 70 seconds. We immediately rinsed the slide with water to remove the excess safranin and after blotting the slide with pieces of paper we examine the smear under the microscope.

upon examination we determine that are bacteria did not contain endospores

Week 4

Week 4, Day 1:
9/17/13

A New Test Tube, A New Chapter

This week, we received a test tube with a new colony of bacteria to work with. Since this bacteria was growing on an agar slant, our first step in identifying the bacteria was to analyze the growth pattern in the test tube. The bacteria we received was growing in what looked like a filiform, or even, pattern.

After observing this growth pattern, we used a sterilized inoculating loop to transfer a small amount of the new bacteria to another test tube. This tube was then placed in the incubator at 37' C to grow over the next few days. 

Using aseptic technique, we then took a loopful of bacteria from the original test tube and mixed it with a drop of distilled water on a new glass slide. We let this mixture air-dry, then passed it through the propane torch three times to heat-fix it, creating a bacterial smear.

Our first step in identifying this bacteria was to determine whether it was gram-positive or gram-negative. To do this, we performed a gram stain on our bacterial smear. We put the slide with the smear on a rack over the sink, covered the smear with crystal violet for 20 seconds, and rinsed the slide with distilled water.

We then covered the smear with Gram's iodine for 1 minute, before rinsing the slide again with distilled water.

After rinsing, we decolorized the smear by dripping 95% ethanol on the slide at a 45' angle until the color from the stain stopped running, then rinsed with distilled water.

We covered the smear with safranin for 1 minute, then rinsed the slide and blotted it with bibulous paper.

By observing the stained smear under the oil immersion lense of our microscope, we saw that our bacteria were gram-negative. Because of their thin peptidoglycan layers, the bacteria had been unable to hold the crystal violet stain, and came out safranin-pink instead.

Week 4, Day 2
9/19/13

After determining last Tuesday that our new bacteria was gram-negative, we spent Thursday's lab preparing negative and capsule stains on both our gram-positive environmental sample bacteria and the gram-negative test tube bacteria we received in class.

To prepare a negative stain on the test tube bacteria, we took two clean slides and placed a small drop of nigrosin at the end of one. Then, using aseptic technique, we used our inoculating loop to transfer a loopful of test tube bacteria to the slide. We then mixed the transferred bacteria with the nigrosin on the slide.

After transferring the bacteria, we took the second (spreader) slide and touched it to the nigrosin/bacteria drop at a 30-45' angle. Once the drop had spread across the length of the spreader slide, we quickly pushed the spreader slide across the first slide, spreading the bacteria/nigrosin mixture across the first slide.

Once this smear had air-dried completely, we examined it under the oil-immersion lense of our microscope. The negative stain allowed us to view the bacteria against a dark background, making it easy to see their shape. This method can also be used to observe bacteria that do not absorb other dyes, or bacteria that are too fragile to undergo heat-fixation when preparing a bacterial smear.

We also repeated this procedure on our environmental sample bacteria. Both bacterial strains appeared to have a spherical shape.

Once we had prepared a negative stain for our environmental and test-tube bacteria samples, we used the negative stains as a foundation for preparing a capsule stain of each type of bacteria. The capsule stain would allow us to view bacterial capsules (the outer coating of a bacteria cell) or slime layers.

To prepare a capsule stain, we took the negative stain slides for the environmental and test-tube bacteria samples, and placed them on a rack over the sink. We covered each slide with safranin for 1 minute, then rinsed off the excess stain with distilled water.

After blotting the slides with bibulous paper, we observed that our bacteria did not have any capsules. However, upon examination, we began to doubt our initial belief that our bacteria was spherical in shape.

We did another bacterial smear and determined that our bacteria was, actually, rod-shaped

Week 3

Week 3

Preparing a Gram Stain. 

9/12/13

This week the whole lab would be working with another isolated colony of their bacterial specimen and observe the colony by means of differential staining as opposed to simple staining. This week, we would focus on preparing and observing a gram stain. The preparation of this stain begins similar to the simple staining we worked on in week 2. We would again find an isolated colony of bacteria from our specimen plate:

We would then place a drop of distilled H2O (water) on a glass slide and proceed to sterilize an inoculating loop.

We used the sterile inoculating loop to capture one of the isolated, bacterial colonies and would transfer it to the drop of water on the slide. Using the inoculating loop, we thinned out the material sample and water across the whole slide.

We then proceeded to follow the directions like good students, and after the slide had air-dried, we "fixed" the smear by passing the glass through a lighted bunsen burner three times.

We then placed the fixed smear on a rack over a lab sink, and covered the smear with a crystal violet stain for 20 seconds

.

After the stain had set on the slide for 20 seconds, we rinsed the slide with distilled water to remove the excess stain. We covered the slide with Gram's iodine for 1 minute and then rinsed the slide again. Next, we held the slide at a 45 degree angle and a 95% ethanol solution (a decolorizing agent) over the slide until the run off was no longer pigmented. Once we achieved this, we rinsed the slide again and then covered it with a safranin dye for 1 minute.

 We then rinsed the slide and blotted it out with a bibulous paper. 

Once the slide was dry, we observed it under a microscope.

The purpose of this experiment was to determine if our bacterial samples were either gram-positive or gram-negative bacteria. Gram-positive bacteria have a thicker peptidogylcan layer than gram-negative bacteria, thus it (in essence) traps the violet and iodine solutions more effectively than the thinner layer found in gram negative bacteria. The 95% ethanol solution dehydrates the the peptidoglycan layer, thinning it out. The thinner layer in gram negative bacteria cannot retain the violet dye, whereas the gram positive bacteria is able to retain the stain. The safranin is added as a counter stain to then stain the now-stainless gram negative bacteria, thus providing us with a differential stain. Here we observed that our bacteria is gram positive, noted by violet stain, as well as spherical shaped. However, we also observed that there was some pinkish dyed bacteria. This could have occurred through heating up the slide too much when fixing the bacterial smear. We would repeat the procedure with a new stain to see if we could correct this error. 

The following lab period we would proceed to finish our second attempt of gram-staining our bacterial smear. We followed the same procedure and, after observation, came up with a more accurate sample. We thereby determined that our bacterial sample was, in fact, gram-positive.