Specimen Collection, Transport and Processing: Bacteriology

F. Pus

  • Purpose: To isolate and identify bacterial etiological agent(s) in deep-seated pus/wound specimens.
  • Specimen collection: Preferably collect specimen prior to initiation of therapy and only from wounds that are clinically infected or deteriorating or that fail to heal over a long period.
  • Cleanse surrounding skin or mucosal surfaces.
  • For closed wounds and aspirates, disinfect with 2% Chlorhexidine or 70% alcohol followed by an iodine solution [1 to 2% tincture of iodine or a 10% solution of povidone-iodine (1% free iodine)]. Remove iodine with alcohol prior to specimen collection.
  • For open wounds, debride, if appropriate, and thoroughly rinse with sterile saline prior to collection. Sample viable infected tissue, rather than superficial debris.
  • Wound or abscess aspirates:
  • Samples collected by using a syringe and needle should be placed in a sterile container or blood collection tube without anticoagulant (g., Vacutainer or similar type) for submission to the laboratory.
  • A portion of the sample should also be placed in a sterile tube containing anaerobic medium like RCM if an anaerobic culture is required.
  • Open wounds:
  • Cleanse the superficial area thoroughly with sterile saline, changing sponges with each application. Remove all superficial exudates.
  • Remove overlying debris with scalpel and swabs or sponges.
  • Collect biopsy or curette sample from base or advancing margin of lesion.
  • Pus:
  • Aspirate the deepest portion of the lesion or exudate with a syringe and needle.
  • Collect a biopsy sample of the advancing margin or base of the infected lesion after excision and drainage.
  • For bite wounds, aspirate pus from the wound, or obtain it at the time of incision, drainage, or debridement of infected wound.
  • Tissues and biopsy samples:
  • Tissue biopsy samples should be collected from areas within and adjacent to the area of infection. Large enough tissue samples should be collected to perform all of the tests required (i.e., 3 to 4 mm biopsy samples).
  • If anaerobic culture is required, a separate piece of tissue should be submitted in a sterile tube containing anaerobic medium like RCM.

Collect swabs only when tissue or aspirate cannot be obtained.

  • Limit swab sampling to wounds that are clinically infected or those that are chronic and non-healing.
  • Remove superficial debris by thorough irrigation and cleansing with non- bacteriostatic sterile saline. If wound is relatively dry, collect with two cotton-tipped swabs moistened with sterile saline.
  • Gently roll swab over the surface of the wound approximately five times, focusing on area where there is evidence of pus or inflamed tissue.

NOTE: Organisms may not be distributed evenly in a burn wound, so sampling different areas of the burn is recommended. Blood cultures should be used to monitor patient status.
Standard precautions to be followed while handling the specimen:
NOTE: Syringes with the needle attached should not be accepted due to the sharps and
Bio hazard risk to staff.

  • Grossly contaminated specimen or leaky containers and collection containers of doubtful sterility must be noted and mentioned.
  • Deliver aspirates and tissues to the laboratory within 30 minutes for best recovery.
  • Keep tissues moist to preserve organism viability.
  • Do not refrigerate or incubate before or during transport. If there is a delay, keep sample at room temperature, because at lower temperature there is likely to be more dissolved oxygen, which could be detrimental to anaerobes.
  • Rejection Criteria:
  • For anaerobic culture, avoid swab collection if aspirates or biopsy samples can be obtained.
  • Do not accept specimens for microbiological analysis in container with formalin.
  • Specimen Processing:                                                                                                             Day 1:
  • Aspirate, pus and swab:
  • Mix the specimen thoroughly. Place a drop of the specimen onto each medium e. RCM, blood agar and MAC.
  • Prepare smear for Gram stain by placing a drop of specimen on a slide and spreading it to make a thin preparation.
  • Perform a Gram stain on all specimens and use in the evaluation of culture
  • Record the relative numbers of WBCs and bacterial and fungal morphotypes. If clinically important organisms are recognized or suspected (g., from a normally sterile site based on the Gram stain interpretation, telephone or report results to the appropriate caregiver immediately
  • Report any bacteria seen in a surgically collected specimen from a normally sterile site.

Aerobic incubation conditions:

  • Incubate RCM, blood agar and MAC in an incubator at 37°C. Incubate for a minimum of 24 hours for open wound cultures. Incubation may be extended to 2-3 days for invasive specimens (e., aspirated fluids and tissues) that remain culture negative after 24-48 hours of aerobic incubation.
  • Critical deep-wounds, abscesses, and tissue samples should have anaerobic cultures requested in order to recover all the primary pathogen(s) causing infection in specific clinical conditions (g., tissue or pus from brain, lung, liver tissue, deep wounds, abscesses, etc.).
  • Reporting results:
  • Report Gram stain results as soon as possible, generally within 1 hour for specimens from critical sites.

DAY 2:

  • Culture interpretation:
  • Report growth on blood agar and McConkey agar.
  • Correlation with results on Gram stain is to be done.
  • In case of no growth on both plates and RCM sterile, report as sterile.
  • In case of just a film of growth on the plates and turbid RCM, plates should be further incubated and subculture from RCM done and looked for growth on the next day.
  • If there is growth of 3 or more organisms on culture plates, report as mixed flora of doubtful significance with suggestion of repeat sample.

The most common urine specimen received is the per-urethral voided urine. Healthy urethra is unsterile and it is extremely critical that urine specimens be collected carefully to minimize Urethral contamination. There are several types of urine specimens and the results of each Types are determined by different guidelines. Therefore, it is essential that each urine specimen Received by the laboratory is clearly labelled as to the type of collection of urine specimen.

  1. Collection of urine:

Midstream clean catch urine:

  • The midstream clean catch urine is the most common type of urine specimen.
  • The technique involved in collection is based on voiding the first portion of urine, which is most likely to be contaminated by urethral commensal.
  • It is recommended that the first voided morning specimen be collected, as bacteria would have multiplied to high levels after overnight incubation in the bladder.
  • If not possible, the urine can be collected during the day, preferably 4 hours after the last void, keeping in mind that the counts may be lower, yet significant.
  • Midstream clean catch urine should be collected in a sterile, wide mouth, screw capped bottle after very thorough preliminary cleaning of external genitalia with soap and water. Antiseptics should not be used for this purpose.
  • Indwelling catheter:
  • Hospitalized patients with indwelling catheter are especially at risk of developing UTI.
  • To avoid contamination, the specimen is collected by disinfecting a portion of the catheter tubing with alcohol & puncturing the tubing directly with a sterile syringe with needle and aspirating the urine.
  • The urine MUST NOT be collected from the drainage bag.
  • Suprapubic collection:
  • The Suprapubic collection is IDEAL and it avoids urethral contamination but is invasive.
  • This procedure is usually reserved for infants and adults, from whom it is difficult to obtain a midstream clean catch urine specimen.
  • Disinfect the skin above the bladder and plunge a sterile needle with syringe into the bladder; aspirate the urine and transfer to a sterile container.
  • Percutaneous nephrostomy (PCN) aspirate:
  • Percutaneous nephrostomy aspirate is urine collected directly from renal pelvis.
  • If the sample is a PCN catheter sample, collection must be done as explained for indwelling catheters and not from the drainage bag.
  • Cystoscopy specimens:
  • Cystoscopy specimen is urine collected from the bladder during Cystoscopy.
  • Ileal conduit specimen:
  • Ileal conduit specimen is collected after cleaning stoma site.
  • A fresh drain of urine is collected. It must not be collected from the urine drainage bag.
  • Intermittent catheter specimen:
  • A red rubber catheter is introduced into the urethra periodically to drain urine from the bladder.
  • It is collected directly into a specimen container.
  1. Specimen Transport:
  • Urine must be transported to the lab as soon as possible.
  • It should be cultured as early as possible after collection, preferably within 2 hours.
  • In case of delay, it may be refrigerated up to a maximum of 24 hours before plating.
  1. Processing of specimen:
  • Smear:
  • Transfer approximately 2 ml of well mixed, un-centrifuged urine specimen using a sterile Pasteur pipette into a labelled tube, and place one-drop of urine on a clean glass slide using the same pipette.
  • Do not spread.
  • Allow to dry (air dry or on a dryer), heat fix and stain by Gram stain.
  • Keep the specimen tubes in the refrigerator till plating and thereafter store the specimen tubes at 2 – 8ºC until the final report is sent.
  • Examination of wet smear of uncentrifuged urine:
  • Look for pus cells and microorganisms.
  • Quantify the presence of pus cells and microorganisms, most commonly Gram-negative bacilli and also Gram-positive cocci, into many, moderate, few or occasional.
  • Also make a note of presence of epithelial cells and other micro-organisms, viz. yeast like organisms, Gram-positive bacilli.
  • Culture:
  • Choice of media and dilution:
  • Results of direct smear examination are used as a guide for choice of media and dilution of specimen as indicated in the Table.

                      Recommended media according to microscopic findings:

       Pus cell Epithelial Cell      Bacteria        Media Recommended
      BA* MA**
           Nil            Nil            Nil ½ plate 0.01 ml undiluted (10 μl) ½ plate 0.01 ml undiluted (10 μl)
(occasional –
           Nil Occasional to few GNB and/ GPC Full plate 0.01 ml undiluted Full plate 0.01 ml undiluted
(occasional –
           Nil Moderate GNB Full plate 0.01 ml of 1/10 diluted urine Full plate 0.01 ml of 1/10 diluted urine
(occasional –
           Nil Moderate to many GNB Full plate 0.01 ml of 1/100 diluted urine Full plate 0.01 ml of 1/100 diluted urine
(occasional –
          Nil    Occasional to many GPC
or YLO or GPB
Full plate 0.01 ml undiluted urine Full plate 0.01 ml undiluted urine
(occasional –
          Nil     Only YLO Full plate n0.01 ml undiluted Full plate 0.01 ml undiluted
add SAB**
Many            Nil No bacteria Full plate 0.01 ml undiluted Full plate 0.01 ml undiluted
(occasional –
    Few to many Many GNB with or without diphtheroids, YLO etc. Full plate
0.01ml of 1/100 diluted urine
+ SAB undiluted urine 0.01ml
Full plate 0.01 ml of 1/100 diluted urine
  • Special situations:
  • Suprapubic collection– use full plate BA and MA
  • PCN aspirate-use full plate BA, MA, CA and thioglycollate broth, irrespective of the smear finding.
  • Make 1:10 dilution when moderate GNB and 1:100 dilutions when many GNB are present in the smear.
  • *4-area streaking without flaming in between for isolation.
  • **Criss-cross streaking for colony count.
  • Inoculation:
  • To evaluate the clinical significance of a growth in urine culture, estimation of the number of organisms present per ml of urine is essential. If needed dilute the urine sample 1:10 or 1:100 using sterile normal saline.
  • For 1:10 dilutions mix 0.5 ml urine with 4.5 ml sterile normal saline.
  • For a 1:100 dilution, mix 0.1 ml (100 μl) urine with 9.9 ml sterile normal saline.
  • Inoculate well-mixed, un-centrifuged, undiluted or diluted urine on to BA and MA using a pipette that delivers 0.01 ml.
  • As shown in the diagram below, streak on BA as guide for 4 area streaking. Be sure to progress from one area to next and DO NOT go over to the previously streaked areas.
  • Do not flame the streaking loop between streak areas. This gives adequate isolation of colonies in the fourth area of streaking.
  • Use a triangular loop on MA to achieve even distribution of the inoculum by rotating the plate (CRISS-CROSS STREAKING). Do NOT flame the loop during this streaking maneuver. If half plate is used on MA, use streaking with streaking loop and spread the inoculum evenly by close streaking, once, over half the plate.
  • If chocolate agar is used, follow the same streaking pattern as for blood agar.
  • Incubate all the inoculated plates aerobically at 37ºC.                                                                                                                                                                               N

Colony Counts:

  • After overnight incubation count the number of colonies manually on each plate and multiply the number of colonies counted by
  • 100 for undiluted urine.
  • 1000 for a 1:10 dilution of urine
  • 10000 for a 1:100 dilution of urine. This gives total number of viable bacteria present in 1.0 ml undiluted urine and express as CFU/ml of urine.
  • Interpretation of Counts:
  • The significance of a positive urine culture is most reliably assessed in terms of the number of colony forming units (viable bacteria) present in the urine. The following is offered as a guide for midstream clean catch urine.
                 <1000 CFU/ml INSIGNIFICANT bacteriuria; UTI-unlikely
1000- 100,000 CFU/ml PROBABLY SIGNIFICANT bacteriuria; UTI probable
                  > 100,000 CFU /ml SIGNIFICANT bacteriuria; UTI certain

For SPC, PCN and cystoscopic specimens, any CFU is significant irrespective of number.

  • Identification of isolates:
  • Identify all coliform bacteria that are considered probably significant.

The preliminary screening media:

  • Mannitol motility medium, triple sugar iron agar medium, peptone water, citrate and if needed include Christensen’s urea agar and lysine iron agar
  • Identify all non-lactose fermenting organisms even if they are in insignificant range in order to rule out Salmonella spp (carrier state).
  • Identify beta haemolytic streptococci.
  • Perform grouping for beta hemolytic streptococci in pregnant women, even if they are few in numbers and along with skin contaminants in order to rule out presence of group B Streptococcus
  • Antimicrobial susceptibility testing:
  • For the panel and detailed methodology of AST please refer to procedure on antimicrobial susceptibility testing.
  • When requested, follow guidelines below for testing organism’s susceptibility to antibiotics.
  • <103 CFU/ml         AST not done, except for cystoscopic, PCN or SPC specimens.
  • 103-105 CFU/ml AST done on two organisms depending on their probable significance and relative numbers
  • >105CFU/ml          AST done on 1-2 organisms and rarely three organisms depending on their significance and relative numbers.
  • Reporting:
  • When there is no growth after 24 hours of incubation, send a preliminary report as “No growth”.
  • When growth shows 1-2 types of organisms with >100,000 CFU/ml (in presence of pus cells), report as “Significant, >100,000 CFU/ml, with organism or organisms”.
  • When growth suggests gross contamination, g. mixture of diphtheroids, Coagulase negative staphylococci, micrococci, YLO and >2 types of GNB and the smear shows pus cells, report as “Mixture of organisms along with contaminants” and suggest repeat “midstream clean-catch” urine sample for culture to confirm significance.

H.Fecal specimens

  • Purpose
  • To describe the collection of faecal samples for microbiological examination, and processing in the laboratory for microbiological examination.
  • Procedure
  • Specimen Collection and Transport
  • A small quantity of solid/semisolid stool or one third of the container in case of watery stool is collected in a sterile screw-capped disposable 40 ml container.
  • A rectal swab is not recommended as the material obtained is never adequate for all the tests or for inoculating all the media used for culture.
  • The sample has to be collected preferably prior to initiation of antibiotics in the container directly, taking care not to soil the outside of the container. Samples should not be collected from bedpan.
  • The sample has to be immediately transported to the laboratory on collection.
  • If there is a delay in transporting faecal specimens or if samples have to be sent by post, one of the following transport media may be employed:
  • Phosphate buffered glycerol saline solution.
  • Stuart’s transport medium.
  • Cary and Blair transport medium.
  • Microscopy
  • For all watery faeces samples, whether the doctor orders or not, examine a hanging drop (HD) immediately, or wet preparation by darkfield (DF) microscopy.
  • If dark-field microscopy is positive, proceed with immobilization test with cholera O1 non-differential and V. cholera O139 specific antiserum and examine again under darkfield microscope.
  • Culture and Isolation
  • Commonly encountered enteric pathogens and potential pathogens include Salmonella, Shigella, cholerae, Arizona, Edwardsiella, Aeromonas, Plesiomonas, diarrhoegenic E. coli and V. parahaemolyticus.
  • Routine media to be included are BA, MA, XLD or DCA, and Selenite F broth.
  • BA is included for all stool samples as primary plating.
  • Use a swab/pasteur pipette.
  • Place a loopful of the specimen over a small area of each plate, then flame the loop, and streak from the inoculated area over the entire plate.
  • Inoculation on DCA: 1 in 10 dilution of specimen in saline is streaked with the help of triangular loop to get maximum isolation of colonies.
  • If cholerae is suspected, a TCBS and alkaline peptone water (APW) medium with a pH 8.4 to 8.6 are also inoculated. After 4 hours of incubation at 37°C, a drop is taken from the surface of the APW is examined under DFM or phase contrast microscopy. A subculture is also made on BA and TCBS.
  • Place 1-2 ml or 1 g of faecal suspension into a tube containing selenite F broth, or pick up an amount approximately the size of a pea and emulsify it in the selenite F broth.
  • For all samples, media incubated are examined after 18 hours and a subculture is done immediately from the selenite F broth onto a DCA plate.
  • Colony characteristics on different media:
  • Deoxycholate citrate agar
  • Shigella species: Opaque ground-glass NLF colonies, with even margins. Salmonella Typhi and Paratyphi: Translucent, colourless NLF colonies.
  • Other Salmonella Species: Large NLF opaque colonies may have a brownish/blackish centre.
  • Vibrio cholera: Small, colourless, translucent colonies, which may appear after 48 hours.
  • Pseudomonas aeruginosa: Large or small NLF colonies, transversely elongated, often with a detectable pigment.
  • Proteus group: Colourless NLF, raised, opaque colonies.
  • coli: Raised, opaque, pink colonies surrounded by a pink halo of precipitated bile salt.
  • Klebsiella and Enterobacter: Mucoid, pink colonies.
  • DCA is examined at the end of 24 hours and again after 48 hours.

Colonies of all non-lactose fermenters (NLF) may look similar: reliance should not be placed on colony characteristics for differentiation. Occasionally 48 hours incubation may be needed for the NLFs to appear.

  • DCA is found to be highly satisfactory for the isolation of non-lactose fermenting faecal pathogens, g. Salmonella and Shigella.
  • The growth of coliform bacteria is inhibited or greatly suppressed. Gram-positive bacteria are generally inhibited.
  • Occasionally, however, coliforms do grow on this medium and such isolates produce acid from lactose, precipitate bile salt, and cause a pink opacity in the medium, which makes it difficult to differentiate the pathogens that may also be present.
  • MacConkey Agar:
  • Shigella species: Colourless NLF colonies varying from small to large and from translucent to moderately opaque.
  • Salmonella Typhi: Colourless NLF colonies, varying from small to large and from translucent to slightly opaque, smooth with even or slightly, irregular leaf-like edges.
  • Other Salmonella species: Colourless NLF colonies, usually more opaque than the above.
  • Vibrio cholera: Medium sized colourless, transparent colonies.
  • Pseudomonas aeruginosa: Large, glistening NLFs, moist colonies, often elongated, with detectable greenish pigment.
  • Proteus group: Large, colourless NLF colonies.
  • coli: Large non-mucoid colonies, with even pink colour.
  • Klebsiella and Enterobacter: Large mucoid colonies, with pink centers and colourless peripheries or even pink.
  • NOTE: Colonies of all non-lactose fermenters (NLF) may look very similar: Reliance should not be placed on colony characteristics for differentiation.
  • Xylose lysine deoxycholate medium
  • Shigella species: Small pink colonies.
  • Salmonella species: Small pink colonies with or without black centers.
  • Coliforms: yellow colonies, or yellow with black centres (Citrobacter freundii and Proteus vulgaris)
  • Pseudomonas: pink colonies.
  • NFGNB: pink colonies
  • Black centre colonies grown on XLD without isolation need to be subcultured on another XLD.
  • Thiosulphate citrate bile salts sucrose (TCBS) agar:
  • cholerae O1 and V. cholerae O139 produce flat yellow disk-like colonies due to the fermentation of sucrose in the medium.
  • parahaemolyticus produces green colonies.
  • Biochemical Tests for Screening:


  • Choose a single colony of each type of isolate from each plate (pink colony from XLD plate) and inoculate the media listed below in that order using a straight needle touching the colony once.
  • Recharge of the needle should not be necessary.
    • Mannitol motility medium: Stab down the centre of the medium, reaching the bottom, but not touching the sides.
    • TSI: Stab into the butt and streak along the surface.
    • Peptone water: Dip the needle into the medium.
    • Citrate utilization test: Inoculate lightly from a young culture over the entire surface of the slant of Simmon’s citrate agar using a straight wire. Incubate at 37ºC for 2 – 7 days.
  • Reading: Blue medium with a streak of growth is positive, g. Klebsiella spp.
  • Original green colour and no growth indicate a negative reaction, g. E.coli
  • LIA: For – black centre colonies on XLD & DCA, stab the butt and streak the slope
  • Examine after overnight incubation.
  • Reading
  • Examine mannitol motility medium for evidence of motility and mannitol fermentation.
  • Examine the TSI for fermentation of glucose, lactose, and sucrose, presence of gas and H2
  • Examine LIA for decarboxylation of lysine and presence of H2S shown as a black precipitation. Note the amount of H2S produced & deamination.
  • Test peptone water culture for the presence of indole after overnight incubation. Indole test after extraction may be done with a lipid solvent like xylol when reactions are doubtful.

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