do a lab summary of minimum 300 words.

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do a lab summary of minimum  300 words.

do a lab summary of minimum 300 words.
23 Motility Some bacterial species have flagella which allow the cells to travel from one place to another. These are referred to as motile species. For the purposes of this lab, we will not deal with other modes of bacterial motility. Those species in our lab devoid of flagella are non-motile. In this lab, we will only consider the motility (qualitatively, i.e. motile or non-motile) of Gram negative rods. Examples of clinically significant non-motile Gram negative rods that will be used in our lab include Klebsiella, Shigella and Acinetobacter. Although Alcaligenes is extremely motile, it only grows very near the top of the agar due to it’s requirement for a lot of oxygen. For this reason, it can be hard to detect Alcaligenes motility. Several methods exist for determining motility, and a few are discussed below. We will use the stabbed soft agar deep method in our lab. Stab a soft agar deep Motility can be determined by monitoring turbidity (cloudiness) following incubation in a stabbed soft agar deep medium such as TSA with 4 – 5gm agar/liter or SIM media (which we will cover in a subsequent lab). Normal hard agar, such as that used for plates or slants, contains 15gm agar/liter. This elevated agar concentration will not allow the movement of any motile cells trapped within the agar matrix. This is not the case with soft agar. To conduct the procedure, cells on the end of a straight needle are stabbed on a straight line into the center of a soft agar deep. Non-motile cells will remain on the stab line as they divide resulting in a distinct line in the agar with no surrounding turbidity. Since soft agar media is slushy motile cells can travel in it. These cells will not remain at the stab line but will radiate outward resulting in turbidity away from a stab line that is visibly less distinct, or even dissipated to invisibility. See image “Motility: left non-motile, right motile.” Highly motile cells can radiate quickly resulting in equal turbidity throughout the media. Less motile cells will radiate, but greater cell density will remain nearer to the stab line. To inoculate a soft agar deep first check your needle to make sure it is straight. Using aseptic technique, collect cells with your needle. Carefully stab the needle through the center of the deep to the bottom of the tube and withdraw along the same path. Do not mix or shake. Incubate at 37oC for 24 to 48 hours, however you cannot incubate motility deeps too long – see point number 2 below. Results from the stabbed soft agar deep method can be misinterpreted. Consider the following points. 1. Soft agar is supposed to be sufficiently oxygen permeable to allow the growth of obligately aerobic species throughout the agar deep. This is not always the case, which can cause confusion when working with obligately aerobic motile cells that have an unusually high oxygen requirement. In this case, look for motility toward the top of the tube. 2. Keep in mind that ANY growth diffusing away from the stab line indicates motility. If not much cell division has occurred then any turbidity present will be difficult to distinguish from the pre-existing translucence of the media. This is perhaps the most common reason for misinterpretation of results using this method. For this reason, eventhough manuals often suggest 24-48 hours of incubation, I tell my students that “when in doubt, incubate longer.” You cannot over-incubate stabbed soft agar deeps. Always use an uninoculated soft agar deep as a control when checking your results. 3. Rarely will a needle follow a laser-straight line to the bottom of the agar deep. The needle will usually follow a crooked path, cutting the agar in one plane or more. Consider a non-motile organism growing along this flat cut path. When viewed perpendicular to the cut plane the growth can easily be mistaken as radiating turbidity. Simply rotate the tube 90 degrees to see if the growth is 3 dimensional (which would indicate actual motility), or if the growth is only following the cut path of the needle. Wet mount / hanging drop method Another way to determine motility is to examine living cells in a wet mount using the hanging drop method. To do this, place a small drop of water in the center of a cover slip using your loop. Collect a small amount of cells with your loop (as if you were making a smear) and gently mix them into the drop of water trying not to spread the drop out. Put small spots of petroleum jelly on the 4 corners of the cover slip. Place a small Sharpie dot next to the drop of water. Lay a depression slide down on the cover slip such that the drop is enclosed in the depression. Press lightly to secure the slip to the slide. Quickly invert the slide so that the drop is hanging from the bottom of the cover slip. View under oil immersion quickly before the heat inactivates the cells or evaporates the drop of water under the slip. The cells will have little contrast and will not be easy to see. Stop down the light with your condenser diaphragm. If everything works correctly you should see motile cells moving around. At first you may confuse the random jerky motion of Brownian motion (caused by random collision of atoms with the cells) for motility. This method is often used to view motile eukaryotic microbes such as protozoa and Euglena, but can also be used to examine the “twiddle and run” motility of bacteria. Cells in the genus Proteus have peritrichous flagella meaning flagella all around the cell, thus they are extremely motile. These cells can even swim on top of agar, a characteristic called “swarming”. For this reason, Proteus can prove very difficult to isolate in a mixed culture, thus causing much frustration to microbiology students. Cells in the genus Pseudomonas have lophotrichous flagella, which is a tuft of flagella on one end of the cell. Lophotrichously flagellated cells are theoretically second only to peritrichously flagellated cells in regard to degree of motility. The remaining “enteric” bacterial genera (Gram negative, oxidasae negative rods) have polar monotrichous flagella, which is a single flagellum on one end of the cell. These are generally considered the least motile cells of the list above, however some strains of polar monotrichously flagellated genera are highly motile. NOTES: * We use SOFT agar deeps for motility tests, NOT tempered hard agar deeps. * Inoculate motility deeps using a needle, NOT a loop. Before inoculating, make needle as straight as possible to limit the diagonal cutting of the agar. * It is hard to over-incubate a motility deep. If in doubt about your results, incubate another day. * Motility results are more easily interpreted while the media is warm (directly out of the incubator). Refrigeration causes the agar to become cloudy. This is a problem. * We will only utilize motility testing no Gram negative organisms this semester. Motility will only be useful in distinguishing the non-motile Gram negative rods (Klebsiella, Shigella and Acinetobacter) from the other Gram negatives. * It is easy to misinterpret a motility result on Pseudomonas due to its obligately aerobic nature. Pseudomonas is motile, but will only show the spreading turbidity toward the top of the tube. * Motility tests should NOT be used to distinguish degrees of motility (quantitatively), but should only be used to distinguish motile from non-motile organisms (qualitatively).
do a lab summary of minimum 300 words.
21 Selective and differential media So far you have used general purpose complex media (such as tryptic soy) to culture a broad range of bacterial species. This media was not formulated to support or inhibit growth of any particular species. We mentioned earlier that samples of mixed cultures on plates could result in confluency (wall-to-wall growth) or TNTC (>300 colonies) if the cell density of the sample is too great. In this case, you could plate a smaller sample or dilute the sample before plating. Another way to decrease colony density is to utilize a selective medium. Selective media are formulated to select for a particular group of (ie. inhibit the growth of all but a particular group of) bacteria. This is especially useful if the organism of interest to you exists in low numbers, and is heavily diluted by the other members of a mixed culture (trying to find a needle in a hay stack). This is a frequent situation in clinical samples. Selective media contain at least one selective agent which inhibits the growth of the unwanted or “contaminating” microorganisms. Selective agents include salts, antibiotics or other inhibitory chemicals at varying concentrations. An example of a selective medium would be TSA with 7.5% added NaCl (table salt). This concentration of salt inhibits the growth of most bacteria, and therefore selects for those that require and/or are capable of growing in the presence of high salt concentrations. The selective media that we will use in this exercise will be Mannitol salt agar and Eosin methylene blue (EMB) agar. Other common selective media include xylose lysine desoxycholate (XLD) agar used for the isolation of Salmonella and Shigella species. Using general purpose complex media, we are able to enumerate the total number of viable cells in a sample by counting the number of colonies that grow up on that media. This is true of pure cultures and mixed cultures alike. What if you wanted to know how many cells of a particular group of bacteria were present in a mixed culture, and you could not depend upon morphology alone to determine which colonies represented that group? One solution would be to plate the sample on a special type of media on which the colonies arising from a particular group of organisms looked unique. Differential media are designed to distinguish certain species, genera or larger physiological groups from others via differences in appearance of colonies, most often color differences of the colonies or surrounding medium. An example of a differential medium would be one containing a particular carbohydrate that could only be fermented to acid by certain microorganisms, mannitol for example. The medium would also contain a pH indicator which would change color as acid accumulated. Those microbes that do not cause the color change must be respiring mannitol, fermenting mannitol to something other than an acid, or using a different carbon and energy source present at a concentration too low to generate enough acid to cause a color change. Another commonly used differential medium is Triple sugar iron (TSI) slants which contains agar, a pH-sensitive dye (phenol red), 1% lactose, 1% sucrose, 0.1% glucose, sodium thiosulfate and ferrous sulfate or ferrous ammonium sulfate. The medium differentiates Gram negative rods in 2 ways. First is by acid production from 1 or more of the carbohydrates. The other is by thiosulfate reduction resulting in the formation H2S, which then interacts with iron to form black FeS. Some media are BOTH selective and differential. Consider the differential mannitol media above with 7.5% NaCl added. You now have a medium that selects for salt tolerant species, and differentiates between those that ferment mannitol to acid vs those that do not. Such a medium exists and is called Mannitol salt agar. Mannitol salt is used to differentiate mannitol fermenting Staphylococcus species (most of which are opportunistic human pathogens) such as S. aureus from non-fermenting Staphylococcus species (most of which are non-pathogens) such as S. epidermidis. See image “Mannitol salt agar.” Micrococcus and Streptococcus are not salt tolerant, and will either fail to grow on Mannitol salt media, or grow poorly. Other commonly used media that are both selective and differential for Gram negative enteric genera include MacConkey’s, Hektoen enteric (HE) and Salmonella Shigella (SS) agars. The differential basis of these media types involves color changes due to acid production from various carbohydrates in the medium. EMB agar is a defined medium which contains lactose as a carbon and energy source. It also contains eosin and methylene blue which inhibit the growth of Gram positive bacteria. The basis for the differential property of EMB media is this: Gram negative lactose fermenting bacteria will take up the purplish-emerald green dyes at a rate proportional to the amount of acid produced from lactose fermentation. This medium will be used in our lab for differentiation of the “enteric” Gram negative rods, otherwise known as the oxidase negative, Gram negative rods. Of the organisms that we are working with this semester, this includes E. coli, E. aerogenes, C. freundii, K. pneumoniae, P. vulgaris, S. enteritidis, and S. flexneri. The term “enteric” implies “of the intestines,” or “of fecal origin.” Appearance of Gram negative, oxidase negative colonies on EMB: The characteristics described below only apply to isolated colonies. As you will see below, much variation exists among the results for most organisms on EMB, depending on crowding, time and temperature of incubation, and variance in media formulation and pH. * Species of the genus Salmonella, Shigella and other non-lactose fermentors commonly produce little or no acid resulting in nearly colorless colonies on EMB. * Enterobacter, Klebsiella and other “Aerogenes-type” organisms commonly produce a moderate amount of acid from lactose fermentation resulting in a characteristic “fish-eye” colony (light with dark center) on EMB agar. Some strains produce more acid resulting in colonies with dark coloration throughout. Alternatively, some cultures of these organisms will form nearly colorless colonies. Although most reports suggest otherwise, certain strains of Klebsiella will produce a transient green metallic-sheen on EMB. See image “EMB agar – fish eye colonies. * E. coli, Citrobacter, Proteus and other “Coli-type” organisms produce relatively large amounts of acid from lactose fermentation resulting in the formation of colonies that are purplish-black throughout, or have a green metallic-sheen. Although the green sheen is most often attributed only to E. coli, the organism given credit as the most robust lactic acid producer, other genera such as Citrobacter and Klebsiella will sometimes form the green sheen. One distinction here is that the green sheen of E. coli is persistent whereas the sheen formed by Klebsiella fades over a period of 6-24 hours. Proteus colonies on EMB are commonly very small and purple-black throughout, although some Proteus cultures form nearly colorless colonies. See image “EMB agar – green metallic sheen. Streak these plates for isolation just as you would streak a TSA plate. Having isolated colonies is not critical for proper results on mannitol salt agar, but IT IS on EMB. You will not see clear fish-eyes, etc. if colonies on EMB agar are not isolated. NOTES * We use mannitol salt agar for Gram positive cocci. * The color change for mannitol salt occurs in the media itself, as well as the colony. * We use EMB agar for Gram negative rods, especially for the oxidase negative variety. * The media that we have discussed in this lab are both selective and differential. Understand these terms. Know the selective and the differential ingredients of each medium discussed. Understand the selective and differential basis of how these work. Know the expected result of each organism discussed above on mannitol salt and EMB.
do a lab summary of minimum 300 words.
18 Colonial morphology: the appearance of a colony If we swabbed the palm of our hand, mix this into a tube of physiological saline or distilled water, streak this onto a plate of media and incubate, many colony types would grow because many different species of bacteria were present in the sample. Some of these colony types would be easily distinguishable, even by the untrained individual because of unique colors (Micrococcus & Pseudomonas), surface luster (Bacillus & Corynebacterium), shape (Proteus), or texture (Klebsiella). For other colony types, the colonial morphology would be quite similar. This is generally true of the Gram negative rods. Colony morphology must be examined closely. With practice, you will learn to see subtle differences between similar colony types. Colony morphology is an important piece of evidence used in identifying organisms, BUT you should never use it as the sole means of making the final call when you have other means of characterization. Remember this when working on your unknown. You should find colony morphology especially helpful in distinguishing the following organisms: *species of the genus Bacillus from other Gram positive rods *Bacillus species one from another *Corynebacterium xerosis from the other Gram positive rods *Staphylococcus species from Streptococcus species *Staphylococcus aureus from the other catalase positive Gram positive cocci *Pseudomonas aeruginosa and Alcaligenes faecalis from the other oxidase positive Gram negative rods *Pseudomonas aeruginosa from Alcaligenes faecalis *Klebsiella pneumoniae from the other oxidase negative Gram negative rods Colony morphology is commonly described using the 7 parameters listed below. You may prefer descriptors other than those listed (ex.: round instead of circular), or you may need to use additional descriptors (such as “terraced” elevation, etc.) if one provided does not accurately describe what you see. Shape: circular irregular rhizoid other? Edge: entire lobate serate other? Density: Opaque translucent transparent Surface: luster Shiny dull wrinkled other? Texture: crumbly hard creamy mucoid other? Elevation: convex flat raised umbonate other? Color: NOTES: * Colony morphology is not very “testable.” The purpose of this exercise is to make you aware of differences in the appearance of colonies so you can use this as another bit of evidence in identification. Images 14 – 38: colony morphology of each organism. See “index of images” on page vii.

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