Objective: To determine the ability of an organism to reduce nitrate to nitrite which is then reduced to free nitrogen gas. The nitrogen in nitrate serves as an electron acceptor. The result of the denitrification process is the production of nitrite:
In this case, all the NO3- will be converted to N2 gas which escapes to the atmosphere. We can test for this step by looking for the absence of NO3- through the addition of Zn powder as described below.
1. Inoculate a nitrate agar slant with your pure culture using a sterile loop to transfer a rather heavy inoculum.
2. Incubate at 37°C for at least 48 hours.
3. Add 2-3 drops of Reagent A and 2-3 drops of Reagent B to your tube. Reagent A is 0.8% sulfanilic acid in 30% acetic acid and Reagent B is 0.6% N,N-dimethyl-α-naphthylamine in 30% acetic acid (CAUTION: Reagent B is a potential carcinogen, so work in the hood and avoid inhaling it or allowing for contact with skin; wash hands thoroughly after work).
Reduction of nitrate to nitrite is indicated if a red color develops quickly (within 1-2 minutes). If no color develops, add a very small amount of zinc powder (~20 mg) to the tube containing the reagents. If a pink to dark red color develops after adding the zinc powder within 5 min., the test is negative (nitrate is present and is not reduced by the organism but zinc has reduced it to nitrite). If no color develops, the test is positive (the organism was able to reduce all the nitrate to nitrite and further to N2 which escaped from the tube).
-- If tubes are stored in the refrigerator, they should first be brought back up to the optimum temperature of the growth condition of the organism.
-- When performing the nitrate reduction test using α-naphthylamine, the color produced in a positive reaction may fade quickly. Interpret results immediately, particularly when performing a number of tests.
-- A strong nitrate-reducing organism may exhibit a brown precipitate immediately after the addition of the reagents. This is due to the effect of excess nitrite upon the p-amino group of the azo-dye and may be reduced by using dimethyl-α-naphthylamine.
-- Some organisms are capable of reducing nitrate to nitrite, yet they destroy the nitrite as fast as it is formed, yielding a false negative result. This nitrite destruction is evident in quite a few bacteria, particularly some Salmonella and Pseudomonas spp. and in Brucella suis.
-- Do not use an excess of zinc; if too much Zn is added, the large amount of hydrogen gas produced may reduce the nitrite (formed from unreduced nitrate) to ammonia (NH3) that could result in a false negative reaction or just a fleeting color reaction.
1. If the plate is refrigerated, it should be allowed to warm up to room temperature and then incubated for 15 min at 37°C before performing the test. Pick a loopful of colonies from a not-too-old pure culture plate and place on a clean glass slide. Do not take your colonies from a blood agar plate. Blood contains catalase; therefore a false positive reaction would be obtained.
2. Add one or two drops of 3% H2O2 and wait 10-15 seconds to observe.
-- Positive test: immediate bubbling (O2 formed).
-- Negative test: no bubbling.
a. When doing the slide test, always add organism to the slide first and then add the reagent since platinum used in the inoculation needle may produce a false positive result. Nichrome wire does not cause bubbling.
b. H2O2 is very unstable when exposed to light. H2O2 decomposition also increases as temperature increases due to dissolved oxygen. Therefore it is important to keep this reagent in the refrigerator at all times when not in use and to shake before it is used.
Objective: To determine the ability of an organism to hydrolyze the glycoside esculin to esculetin and glucose. Esculetin reacts with iron to form a dark brown to black complex. The medium contains 40% bile. Some streptococci that are capable of splitting esculin cannot tolerate an increased concentration of bile. So this is basically two tests: (a) growth on 40% bile and (b) hydrolysis of esculin.
Obtain a bile esculin slant. With a sterile loop, touch a colony of your pure culture to obtain a light inoculum. Uncap the tube and flame the lip of the tube. Insert the loop to the bottom surface of the agar. Touch the agar and gently slide the loop in a zigzag fashion along the surface of the agar as you pull the loop out. Flame the lip of the tube again and put the cap back on. Incubate the slant at 37°C for 48 hrs and check the results. If the results seem negative, continue incubation for up to 96 hrs before reporting the results as negative.
-- Positive: Half or more of the medium is blackened (black to dark brown) in any time interval.
-- Negative: Less than half of the tube is blackened after 96 hrs.
Objective: To determine the ability of an organism to produce proteolytic-like enzymes (gelatinases) which break down gelatin. Gelatinase destroys (hydrolyzes) the gel and causes its collapse and liquefaction.
1. Obtain two solidified gelatin butts but keep them in the refrigerator until just prior to inoculation. Pick up a heavy inoculum from your pure culture and stab one of the butts to a depth of 2 inches. The other tube should not be inoculated and used as a control.
2. Incubate both the test and control tubes simultaneously at the optimal growth temperature for the organism for 48 hours to 14 days.
3. At the end of each 48-hour period, place both tubes (test bacterium and control) in a refrigerator for about 1 hour to determine whether digestion of gelatin (liquefaction) has occurred. Make the transfer from incubator to refrigerator without shaking the tubes. Check tubes daily up to 2 weeks unless liquefaction occurs sooner.
Gently tilt the tubes. The test is positive if the medium of the test organism is liquefied (gelatin breakdown) while that of the control has remained solid (lack of gelatin hydrolysis). The result of the test is negative if the medium of the test organism is as solid as that of the control.
-- Always run a control tube in parallel with organism being tested.
-- Gelatin is solid when incubated at 20°C or less and liquid at 30°C or greater. Gelatin changes from a gel (solid state) to a liquid at about 28°C. Therefore, if gelatin tubes are incubated at 30°C or greater, they must first be placed in a refrigerator for an hour and cooled before an interpretation of liquefaction is made.
-- Do not shake gelatin tubes while warm since growth and liquefaction of gelatin frequently occur only on the surface layer. If the gelatin is shaken and allowed to be mixed with the warm fluid of the medium, there is a possibility that a positive result may be overlooked, and thereby a false negative result may be obtained.
The first fluorescence-based flow cytometry device (ICP 11) was developed in 1968 by Wolfgang Göhde from the University of Münster, Germany and first commercialized in 1968/69 by German developer and manufacturer Partec through Phywe AG in Göttingen. At that time, absorption methods were still widely favored by other scientists over fluorescence methods. The original name of the flow cytometry technology was pulse cytophotometry (German: Impulszytophotometrie). Only 10 years later in 1978, at the Conference of the American Engineering Foundation in Pensacola, Florida, the name was changed to flow cytometry, a term that quickly became popular. Soon after, flow cytometry instruments were developed, including the Cytofluorograph (1971) from Bio/Physics Systems Inc. (later: Ortho Diagnostics), the PAS 8000 (1973) from Partec, the first FACS instrument from Becton Dickinson (1974), the ICP 22 (1975) from Partec/Phywe and the Epics from Coulter (1977/78).
Principle of flow cytometry
A beam of light (usually laser light) of a single wavelength is directed onto a hydrodynamically-focused stream of fluid. A number of detectors are aimed at the point where the stream passes through the light beam: one in line with the light beam (Forward Scatter or FSC) and several perpendicular to it (Side Scatter (SSC) and one or more fluorescent detectors).
Each suspended particle from 0.2 to 150 micrometers passing through the beam scatters the light in some way, and fluorescent chemicals found in the particle or attached to the particle may be excited into emitting light at a longer wavelength than the light source. This combination of scattered and fluorescent light is picked up by the detectors, and, by analyzing fluctuations in brightness at each detector (one for each fluorescent emission peak), it is then possible to derive various types of information about the physical and chemical structure of each individual particle.
Learning the art of ECG interpretation requires intellect, commitment, effort and perhaps most importantly...an organized approach. I personally have spent thousands of hours (yes thousands) looking at 12-lead ECG tracings, studying ECGs for the cardiology boards, interpreting ECGs for direct patient care and developing the ECG tutorials and quizzes of LearnTheHeart.com.
I assume that most of you reading this blog do not have that much time...so let me share with you what I have discovered in my years teaching ECGs to make the process more simple and perhaps even enjoyable.
Read more: 10 Steps to Learn ECG Interpretation
How will samples required for PCR, ELISA, or other immunoassay applications affect which pipette should be used?
• Thick blood smear
• Concentration techniques: mem-brane filtration, microhematocrit centrifugation, lysed venous blood technique, lysed capillary blood technique.
A thick blood smear is spread from 20 μl of capillary blood on a glass slide, air-dried, and stained with a Romanowsky stain. If microfilariae are not detected in thick smears prepared from capillary blood collected at the appropriate time, and if clinical suspicion is strong, concentration techniques are employed. This is because circulating microfilariae are often scanty and sensitivity of microfilarial detection increases when volume of blood sampled is increased.
Membrane filtration: This is a sensitive method but is expensive for routine use in endemic areas. Anticoagulated venous blood (10 ml) is passed through a polycarbonate membrane filter of 3 μm or 5 μm pore size. Following this 10 ml of methylene blue saline solution is passed through the filter for staining the microfilariae. Microfilariae are trapped and retained on the filter, which is placed on a glass slide and examined under the microscope.
Microhematocrit tube or capillary tube method: Two heparinised capillary tubes are filled with blood from skin punctures (or two plain capillary tubes are filled with anticoagulated venous blood). After sealing the dry ends with a suitable sealant, tubes are centrifuged in a microhematocrit centrifuge for about 5 minutes. The capillary tubes are placed on a glass slide and fixed with adhesive tape. Plasma just above the buffy coat layer is examined for motile microfilariae under the microscope.
Lysed venous blood method: 10 ml of venous blood is lysed by saponin-saline solution. The hemolysate is centrifuged, supernatant is discarded, and the sediment is placed on glass slide. After adding a drop of methylene blue solution, a coverslip is placed, and the preparation is examined under the microscope for microfilariae.
Lysed capillary blood method: 0.1 ml of blood obtained by skin puncture is added to 1 ml of saponin-saline solution to cause lysis of red cells. After centrifugation, supernatant is discarded and sediment is placed on a glass slide. A drop of methylene blue solution is added and a coverslip is placed over it. The entire preparation is examined under the microscope for motile microfilariae.
Morphology of Microfilariae on Romanowsky-stained Blood Smears Wuchereria bancrofti: Microfilariae measure about 300 μ in length and 8 μ in breadth. They have a hyaline sheath, which stains pink. There are distinct nuclei in the central axis of the body. Nuclei are not present in the tip of the tail. Cephalic space (present at the anterior end) is as long as it is broad. Tip of the tail is bent backwards and body curves are few.
Brugia malayi: These measure about 230 μ × 6 μ in size. Sheath stains dark pink in color. The nuclei are crowded in the body, are blurred in outline, and the tip of the tail shows two distinct nuclei. Cephalic space is twice as long as it is broad. Instead of smooth curves to the body, there are kinks.
- Spectrophotometer or photoelectric colorimeter
- Pipette 5 ml
- Sahli’s pipette
- Drabkin’s Solution
- Cyanmethemoglobin standard solution with known hemoglobin value
- Take 5 ml of Drabkin’s solution in a test tube and add 20 μl of blood. This way, we will get the dilution of 1:25. Now mix the mixture and allow to stand for atleast 5 minutes. This time is adequate for transformation of hemoglobin to hemiglobincyanide.
- Pour the test sample to a cuvette and read the absorbance of the test sample in a spectrophotometer at 540 nanometer or in a photoelectric colorimeter using a yellow-green filter. Also read the absorbance of the standard solution. Absorbance must be read against Drabkin’s solution.
- From the formula given below, the hemoglobin value is derived.
Preparation of table and graph: Result can be obtained quickly, if the table of graph is prepared which correspond absorbance with hemoglobin concentration. This is markedly acceptable when huge number of samples are daily processed on the same instrument.
- The hemiglobincyanide solution is stable so that delay in getting the reading of absorbance does not influence the result.
- High TLC (total leukocyte cunt) (> 25,000/μl), abnormal plasma proteins (e.g. in Waldenström’s macroglobulinemia, multiple myeloma) or lipemic blood (hypertriglyceridemia), can cause the error in results.