Relevant clinical information is needed to help the laboratory in processing the specimen optimally. The attending doctor’s name and contact number should be clearly indicated, as he may need to be contacted for urgent or critical results.
Recent, current or intended antimicrobial treatment should be indicated to help the microbiologist interpret test results and to help in the selection of antimicrobials to be tested. Previous culture or serological results should be given where possible. The microbiologist should first be consulted if the attending clinician is unsure of which appropriate test(s) to request for.
(a) to ensure the survival and isolation of fastidious organisms and to prevent overgrowth by more hardy bacteria;
(b) to shorten the duration of specimen contact with some local anaesthetics used during collection that may have antibacterial activity.
(a) Put on a pair of sterile gloves.
(b) Wipe the diaphragm of the culture bottle with 70% alcohol (do not use iodine). Cleanse and disinfect intact skin as detailed above in the section on Sample Collection (Collection, point 4).
(c) Allow sufficient standing time for the disinfectant to dry. Do not palpate the vein after disinfecting the skin prior to inserting needle. (Gowns and sterile drapes are not needed if the above is strictly followed.)
(d) Draw blood through a needle and inoculate blood culture bottles. It is not necessary to change the needle prior to inoculation of the bottles in order to minimise the risk of a needlestick injury. If the blood drawn is to be also used for other investigations, e.g. electrolytes, inoculate the blood culture bottles first.
(e) If iodine was used in skin preparation, wipe off residual iodine from patient’s skin with 70% alcohol to prevent skin irritation.
(f) Do not refrigerate the bottles. If delay in transport to the laboratory is anticipated, you may leave the bottles at room temperature.
(g) The BACTEC bottles in use are: Aerobic Plus/F for aerobic bacterial blood culture, Anaerobic Plus/F for anaerobic bacterial blood culture and Myco F/Lytic for fungal blood culture. Do not paste labels over the bar codes of the bottles.
(a) Each blood culture set may consist of an aerobic and an anaerobic bottle.
(b) Usually, two or three culture sets drawn from different sites should suffice. Do not send only one culture set as intermittent bacteraemia may be missed and it may be difficult to determine the significance of certain bacteria. Try not to draw blood through indwelling intravascular catheters.
(c) Recommended volumes : Bacterial culture : 8 to 10 ml of blood from adult patients should be inoculated in each bottle. Fungal culture : 1 to 5 ml of blood should be inoculated into each bottle. Pediatric patients : 1 to 3 ml of blood in each bottle, total volume of blood cultured should not exceed 1% of estimated total blood volume.
(d) Timing of blood cultures (regular intervals or in relation to fever) is not as important as the total blood volume cultured.
(e) Preferably, obtain cultures before the use of systemic antimicrobials.
(f) Acute sepsis, meningitis, osteomyelitis, arthritis, pneumonia: obtain two or three sets of blood cultures.
(g) Fever of unknown origin (such as that caused by an occult abscess): obtain two sets initially, then another two sets 24 - 36 hours later.
(h) Suspected early typhoid fever or brucellosis: obtain four sets over 24 - 36 hours.
(i) Infective endocarditis:
Acute, no antibiotics given within the past two weeks : Blood cultures should be drawn immediately to avoid unnecessary delays in treatment. Obtain three blood culture sets within a 30-minute period before administration of empiric antimicrobial agents.
Subacute, or antibiotics given within past two weeks, or prosthetic valve: Obtain three blood culture sets, spaced 30 minutes to 1 hour apart. This may help document a continuous bacteraemia. If initial cultures are negative at 24 hours, obtain two more sets, for a total of five sets overall.
Suggested volumes are 1 and 2 mL for bacterial and fungal direct smears, cultures and antigen testing respectively. Please note that CSF antigen detection (for S. pneumoniae antigen) must always be ordered concurrently with a CSF bacterial culture, with sufficient specimen for both tests.
(a) Lumbar puncture
(i) Clean the puncture site with 70% alcohol and 10% povidone-iodine or 1 – 2% tincture of iodine before needle insertion.
(ii) Insert the needle with stylet at the L3-L4, L4-L5 or L5-S1 interspace. When the subarachnoid space is reached, remove the stylet and spinal fluid will appear in the needle hub.
(iii) Slowly drain the CSF into the sterile leak-proof tubes. Three tubes are generally required for microbiology, haematology and biochemistry testing. Send the second tube for microbiology and the third tube for haematology. In any case, always send the most turbid tube to microbiology.
(b) Ommaya reservoir fluid
(i) Clean the Ommaya reservoir with antiseptic solution and alcohol.
(ii) Remove Ommaya fluid via the Ommaya reservoir unit, and place it in a sterile tube.
(a) Brain abscess
(i) Aspirate material from the lesion and send it immediately to the laboratory in an anaerobic transport medium, or the syringe, or a sterile container.
(ii) Request for Gram stain and culture (aerobic and anaerobic).
(b) CNS biopsy
Send sample obtained at surgery in a sterile container or in an anaerobic transport medium. Do not add formalin.
Submitted primarily for the detection of Salmonella, Shigella, Campylobacter and Vibrio and in certain cases, Yersinia, enteropathogenic E. coli and Clostridium difficile. For enteric pathogens other than C. difficile, do not send more than 3 stool specimens and not after the third hospital day.
(a) Do not use toilet paper to collect stool. Toilet paper may be impregnated with barium salts which are inhibitory for some faecal pathogens.
(b) Transport stool to the laboratory within two hours.
(c) Obtain stool by having the patient pass stool into a clean, dry bedpan and transfer stool into the container.
(d) Rectal swab
In general, rectal swab specimens have lower sensitivity than stool specimens for fecal pathogens. Their major utility is in screening for Vancomycin Resistant Enterococcus (VRE) and Carbapenemase-producing Enterobacterales (CPE, formerly referred to as CRE). To perform a rectal swab, pass the tip of a sterile swab approximately 2.5 cm beyond the anal sphincter. Carefully rotate the swab to sample the anal crypts and withdraw the swab. The swab must be heavily loaded with faeces. For detection of N. gonorrhoeae, plate immediately on GC-lect medium obtainable from Client Services (Tel: 6321 4950/4952/4904).
Submitted primarily for the detection of lower respiratory tract pathogens in patients (most frequently children) unable to produce quality sputum. Should be performed after the patient wakes in the morning, so that sputum swallowed during sleep is still in the stomach.
Pass a well-lubricated tube orally or nasally through to the stomach of the patient, and perform the lavage. Before removing the tube, release the suction and clamp to prevent mucosal trauma or aspiration (see sub-section in Mycobacteriology).
Submitted primarily for the detection of Giardia and larvae of Strongyloides stercoralis and Ascaris lumbricoides. Pass a tube orally through to the duodenum of the patient. To aspirate a sample for giardiasis, the end of the tube should be at least in the third portion of the duodenum.
This is for detection of Helicobacter pylori in the stomach and duodenum, and Strongyloides stercoralis and Giardia lamblia in the duodenum. Keep the sample moist in a small amount of sterile saline in a sterile bottle, and despatch to the laboratory without delay. Please transport specimens for Helicobacter pylori culture in ice.
Small bowel biopsy should also be sent for histopathological examination if you are looking for Giardia, Cryptosporidium, Microsporidium and Helicobacter pylori.
Swab perianal area when patient gets up in the morning before patient bathes or defaecates. You may use a commercial collection kit consisting of sticky cellophane tape.
For the detection of Neisseria gonorrhoea, Chlamydia trachomatis, Mycoplasma hominis and Ureaplasma sp. and Haemophilus ducreyi, see instructions under the respective headings in "Test Listing”.
For the detection of Herpes simplex virus, see sub-section on Virology.
(a) Amniotic fluid
Aspirate fluid by catheter, at Caesarean section or at amniocentesis.
(b) Bartholin gland
Decontaminate the skin with antiseptic and aspirate material from the duct(s).
Do not use lubricant during procedure as certain lubricants may have an inhibitory effect on microorganisms. Wipe the cervix clean of the vaginal secretion and mucus. Insert a sterile swab stick into the endocervical canal and rotate the swab. Allow the swab to remain in place for a few seconds and remove it.
Collect endometrium samples by transcervical aspiration through a telescoping catheter.
(e) Fallopian tubes
Obtain aspirates (preferably) or swab samples during surgery. Bronchoscopy cytology brushes may be used if exudate is not expressed.
Collect specimens one hour or more after patient has urinated. Stimulate discharge by gently massaging the urethra against the pubic symphysis through the vagina. Collect the discharge with a sterile swab. If discharge cannot be obtained, wash external urethra with soap and rinse with water. Insert a urethrogenital swab 2 to 4 cm into the endourethra, gently rotate the swab, and leave it in place for 1 to 2 seconds. Withdraw the swab and submit it for culture using the appropriate transport media for Neisseria gonorrhoea.
Use a speculum without lubricant in collecting vaginal specimens.
High vaginal swab: Collect secretions from the mucosal wall high in the vaginal canal (near the cervical area) with sterile pipette or swab. In general, a high vaginal swab is not useful for investigating the cause of a pelvic infection, although it is useful for vaginal infection. Vaginitis is most commonly due to Trichomoniasis, Candidiasis or Bacterial Vaginosis.
Low vaginal swab: Insert sterile swab 1 to 2 cm into the lower entrance of the vagina and collect secretions. For determining Group B Streptococcal (GBS) carriage, a low vaginal swab should be taken and sent in suitable transport medium (e.g. using Transwab® with semisolid modified Amies medium). Screening for GBS carriage is best accomplished by a combination of low vaginal and rectal swab/s. Swab the lower vagina (vaginal introitus), followed by the rectum using the same swab or two different swabs. Swabbing both the lower vagina and rectum increases the culture yield compared with sampling the cervix of the vagina without also swabbing the rectum.
Penicillin remains the drug of choice for Group B Streptococcus in non-allergic patients, and no routine susceptibility testing is necessary for vaginal swab isolates. However, if a female patient being screened for Group B Streptococcal carriage is allergic to penicillin or other beta-lactams, please indicate this clearly in the test request. This will prompt the laboratory to test any Group B Streptococcus isolate(s) for susceptibility to alternative agents such as clindamycin and vancomycin.
(a) Anal swab for gonococcal infection and HSV – see instructions given in “Neisseria gonorrhoeae (Gonococcus) Culture” and sub-sections on Virology respectively.
Use a needle and syringe to aspirate material from the epididymis after skin disinfection.
(c) Penile lesion
Clean the surface of the lesion with 0.85% NaCl. Remove any crust if necessary. Scrape the lesion until serous fluid emerges. For HSV, see instructions given in sub-section on Virology. For Haemophilus ducreyi (Chancroid), see instructions given in “Haemophilus ducreyi (Chancroid) Culture”.
(d) Prostatic specimens
For prostatic massage urine specimens, see “Prostatic massage” under “Urine” at the end of this page. A prostatic abscess detected by ultrasound may be aspirated. Prostatic chips may be sent in a sterile container.
For N. gonorrhoeae, see instruction given in “Neisseria gonorrhoeae (Gonococcus) Culture”.
(a) 1 or 2 drops of topical anaesthetic are generally instilled.
(b) Scrape the lower tarsal conjunctiva with a sterilised Kimura spatula.
(c) Inoculate the appropriate media directly (blood and chocolate plate for bacteria, GC-lect plate for Neisseria gonorrhoeae).
(d) Prepare smears by applying the scraping in a circular manner to a clean glass slide or by compressing material between two glass slides.
(a) 1 or 2 drops of topical anaesthetic are generally instilled.
(b) Using short, firm strokes in one direction, scrape multiple areas of ulceration and suppuration with a sterilised Kimura spatula.
(c) Inoculate each scraping directly onto appropriate media.
(d) Prepare smears by applying the scrapings in a gentle circular motion over a clean glass slide or by compressing material between two glass slides.
(a) Samples are obtained by needle aspiration or vitrectomy.
(b) Inoculate appropriate solid or liquid culture media directly (do not try plating more than 0.5 mL of fluid on agar plate) or transport the sample in a capped sterile syringe (without the needle) or place larger volumes of vitreous material into a sterile container. Transport to the laboratory immediately.
(a) 24-hour sputum collections are not recommended for culture.
(b) Sputum cultures for usual bacterial culture are less likely to be meaningful if the patient has been on antibiotics, or if there is a dry non-productive cough.
(c) If Corynebacterium diphtheriae is suspected, inform the laboratory beforehand, so that the appropriate media may be prepared. For Neisseria gonorrhoeae and specimens from neonates for Chlamydia trachomatis and Mycoplasma hominis and Ureaplasma sp. detection, contact Client Services (Tel: 6321 4950/4952/4904) for transport media or Teflon slide.
(d) For microscopy for Pneumocystis jirovecii (formerly P. carinii), bronchoalveolar lavage (BAL) should be sent. Induced sputum with 3% hypertonic saline may be obtained but the procedure should be done by skilled personnel trained in the technique (see below).
(a) Expectorated sputum
(i) If possible, have the patient rinse mouth and gargle with water prior to sputum collection.
(ii) Instruct the patient not to expectorate saliva or post-nasal discharge into the container.
(iii) Collect sample resulting from deep cough in sterile screw-capped container.
(b) Induced sputum
(i) Using a wet toothbrush, brush the buccal mucosa, tongue and gums before the procedure.
(ii) Rinse the patient’s mouth thoroughly with water.
(iii) Using an ultrasonic nebulizer, have the patient inhale approximately 20 to 30 mL of 3 to 10% NaCl.
(iv) Collect the induced sputum in a sterile screw-capped container.
(c) Tracheostomy aspiration
Aspirate the specimen into a sterile sputum trap. Tracheostomy is followed by colonisation within 24 hours of insertion of the tube. Results must be correlated with clinical and X-ray findings.
(d) Endotracheal aspirate
Suction the excess secretions in the mouth around the tube. Using a new suction catheter, place it within the tube and aspirate into a sterile sputum trap. Endotracheal tubes (ETT) are often colonised with bacteria and results should be correlated with clinical and X-ray findings. ETT tips are not acceptable samples.
(e) Bronchoscopy samples
These include bronchoalveolar lavage (BAL), bronchial washing, bronchial brushing and transbronchial biopsy samples. For bacterial culture, BAL or bronchial brushing using a protected bronchial brush is preferred.
(i) Pass the bronchoscope transnasally or transorally in non-intubated patients or via the endotracheal tube in intubated patients.
(ii) Wedge the tip of the bronchoscope in a segmental (for bronchial wash) or subsegmental (for BAL) bronchus.
(iii) To obtain specimens for bronchial wash or BAL, inject sterile nonbacteriostatic 0.85% NaCl (generally 5 to 20 mL aliquots) from a syringe through a biopsy channel of the bronchoscope. Gently suction the 0.85% NaCl into a sterile container before administering the next aliquot. (In general, 50 – 75% of the 0.85% NaCl instilled is recovered in the lavage effluent). Keep aliquots separate during collection. Combine aliquots from the same site for microbiology cultures and smears
(iv) Bronchial brush specimens
Insert a telescoping double catheter plugged with polyethylene glycol at the distal end (to prevent contamination of the bronchial brush) through the biopsy channel of the bronchoscope.
(v) Transbronchial biopsy
Obtain the biopsy sample through the biopsy channel of the bronchoscope, and transport it in a sterile container with a small amount of sterile non-bacteriostatic 0.85% NaCl.
(f) Lung aspiration
Using CT Scan guidance, obtain lung aspirate by inserting a needle through the chest wall into a pulmonary infiltrate. Aspirate material from the lesion. If the lesion is large or if there are multiple lesions, collect multiple specimens from representative sites. Send the aspirate in a sterile container and/or anaerobic transport medium.
(g) Lung biopsy
Obtain a 1 to 3 cm square piece of tissue if possible. If the lesion is large or if there are multiple lesions, collect multiple specimens from representative sites. Submit in a sterile container(s) without formalin.
(h) Pleural fluid
(i) Clean the puncture site with 70% alcohol and iodine solution (1% tincture of iodine or 10% povidone-iodine). Aspirate and send as much pleural fluid as possible in a sterile container and/or in anaerobic transport medium. Taking samples from drains is not encouraged as any growth may represent colonization. Please indicate in the test request if this has been done.
(ii) Request for Gram stain and culture (aerobic and anaerobic).
Submitted primarily for detection of group A streptococcus. For pharyngeal infection with N. gonorrhoeae, plate on GC-lect medium, see instruction given in ”Neisseria gonorrhoeae (Gonococcus) Culture”.
(i) Depress tongue gently with tongue depressor.
(ii) Extend sterile swab between the tonsillar pillars and behind the uvula. Avoid touching the cheeks, tongue, uvula or lips.
(iii) Sweep the swab back and forth across the posterior pharynx, tonsillar areas and any inflamed or ulcerated area to obtain sample.
Submitted primarily for the detection of MRSA carriers.
(i) Insert a sterile swab into the nose. Rotate the swab against the anterior nares.
(ii) Repeat the process on the other side with the same swab.
Submitted for the detection of carriers of group A Streptococcus, N. meningitidis, C. diphtheriae and B. pertussis. Suction material from the nasopharynx and collect it in a sterile container.
(i) Using a syringe aspiration technique, obtain material from the maxillary, frontal or other sinuses.
(ii) Send the sample in the syringe or a sterile container.
Submitted primarily to diagnose middle ear infections only if previous therapy has failed.
(i) Clean the external canal with mild detergent.
(ii) Using a syringe aspiration technique, obtain fluid through the ear drum. Send the sample in the syringe (capped and without the needle) or in a sterile container.
(iii) If the ear drum is ruptured, collect exudate by inserting a sterile swab through an auditory speculum.
For fungal infections of the lung, lung biopsy or aspirates are the best specimens. Otherwise, collect three early-morning fresh specimens of 3 – 5 mL each from deep cough or sputum induction.
Anaerobes should only be sought from sinus aspirates, tympanocentesis fluid, lung aspirates, protected BAL, protected specimen brushings and biopsy specimens. Sputum and bronchial washings are not acceptable as they are usually contaminated with oral anaerobes.
Surface swabs may reflect colonising organisms rather than invasive species. If necessary, try to obtain tissue from an infected area at the margins where bacteria are invading healthy tissue.
(a) Use sterile normal saline to clean or wash off any surface debris (this will also remove colonising flora).
(b) Blot off excess saline with sterile gauze.
(c) Use sterile swab and rotate the tip of swab over the open wound. A swab with transport media should be used, e.g. eSwab with Amies medium.
(d) Superficial wound specimens are not appropriate for anaerobic culture.
(a) Clean the surface with sterile saline.
(b) Using a scalpel blade, scrape the periphery of the lesion. Samples from scalp lesions should include hair that is selectively collected for examination. If there is nail involvement, obtain scrapings of debris or material beneath the nail plate. Collect into sterile container. Skin scrapings can be collected in a sterile container or sandwiched between 2 slides (precleaned with 70% alcohol) and taped together.
(a) Clean the area with sterile saline.
(b) Remove overlying debris.
(c) Curette the base of the ulcer or nodule.
(d) If exudate is present from ulcer or nodule, collect it with a syringe or sterile swab.
Aspirate pus from the wound, or obtain it at the time of incision, drainage or debridement of infected wound. Do not culture fresh bite or trauma wounds as relevant infectious agents will likely not be recovered. Indicate clearly on the request form if this is a bite wound and the animal involved if relevant, so that the laboratory can look for specific pathogens.
(a) Obtain bone specimen at surgery.
(b) Submit in sterile container without formalin. Sample may be kept moist with sterile 0.85% NaCl.
(a) Disinfect the surface with 70% alcohol and then with an iodine solution (1 to 2% tincture of iodine or 10% povidone-iodine).
(b) Aspirate the deepest portion of the lesion, avoiding wound surface contamination. If collection is done at surgery, a portion of the abscess wall should also be sent for culture.
(c) For abdominal and other deep abscesses where anaerobic infection is likely, send for Gram stain and culture (aerobic and anaerobic).
(a) Disinfect the surface with 70% alcohol and then with an iodine solution (1 to 2% tincture of iodine or 10% povidone-iodine).
(b) Aspirate the deepest portion of the lesion or sinus tract. Be careful to avoid wound surface contamination.
(a) Biopsy specimens or aspirates are better than swab specimens.
(b) If vesicle is present, collect both fluid and cells from the base of the lesion.
(c) For actinomycosis, send pus immediately in syringe, sterile container or anaerobic transport medium.
Do not force fluids to make the patient void urine. Excessive fluid intake will dilute and may decrease the organism count
(a) Never collect urine from a bedpan or urinal.
(b) Thoroughly clean the urethral opening (and vaginal vestibule in females) prior to collection procedures to ensure that the specimen obtained is not contaminated with colonizing micro-organisms in this area.
(c) Soap rather than disinfectants, is recommended for cleaning the urethral area. If disinfectants are introduced into the urine during collection, they may be inhibitory to the growth of the microorganisms.
(d) Transport specimen to the laboratory within two hours of collection. If a delay is expected, urine can be stored in the refrigerator at 4°C for a maximum of 24 hours.
(e) Use sterile containers to transport urine. Alternatively, use a dip-slide for urine culture if urine is collected after office hours, on weekends or public holidays.
(f) Any urine collection involving catheterisation should be done with scrupulous aseptic technique to avoid introducing microorganisms into the bladder. Catheterisation solely for the purpose of obtaining a urine specimen for culture is not usually recommended.
(g) Send the first morning voided urine.
(h) Do not submit 24-hour urine collections for culture.
(i) Foley catheter tips are also not suitable samples.
(i) The person collecting the urine should wash hands with soap and water, rinse and dry. If the patient is collecting the sample, she should be given detailed instructions.
(ii) Cleanse the urethral opening and vaginal vestibular area with soapy water or clean gauze pads soaked with liquid soap, from front-to-back.
(iii) Rinse the area well with water or wet gauze wipes, with the same front-to-back motion. Use each pad/wipe only once.
(iv) Hold labia apart during voiding.
(v) Allow a few mL of urine to pass. Do not stop the flow of urine.
(vi) Collect the midstream portion of urine in a sterile container.
(i) The person collecting the urine should wash hands with soap and water, rinse and dry. If the patient is collecting the sample, he should be given detailed instructions.
(ii) Cleanse the penis, retract the foreskin (if not circumcised), and wash with soapy water.
(iii) Rinse the area well with sterile water.
(iv) Keeping the foreskin retracted, allow a few mL of urine to pass. Do not stop the flow of urine.
(v) Collect the midstream portion of urine in a sterile container.
(i) Check dip-slide to make sure media has not fallen off, been contaminated or dried up.
(ii) Collect mid-stream urine in another sterile container. Do not use the dip-slide container to collect the mid-stream urine.
(iii) Dip the slide into the urine for 2 seconds. There must be sufficient urine to cover all the media on the slide.
(iv) Drain off the excess urine from the slide.
(v) Replace the dip-slide in its own sterile container and screw the cap into place.
(i) Remove the external urinary appliance and discard the urine within the appliance.
(ii) Gently swab and clean the stomal opening with a 70% alcohol pad and then with an iodine solution (1 to 2% tincture of iodine or 10% povidone-iodine). Using sterile technique, insert a double catheter into the stoma. (A double catheter helps to minimise contamination of the sample with skin flora).
(iii) Catheterise the ileal conduit to a depth beyond the fascial level.
(iv) Collect the urine drained into a sterile container.
Insertion of a catheter solely for the purpose of collecting a urine specimen is generally discouraged. It may be considered when clean-catch urine is unobtainable and a diagnosis is critical. Indicate on the test request that the specimen was obtained by in/out catheterisation.
(i) Clean the patient’s urethral opening (and, in females, the vaginal vestibule) with soap, and carefully rinse the area with water.
(ii) Using aseptic technique, pass a catheter into the bladder.
(iii) Discard the initial 15 – 30 mL of urine.
(iv) Collect a sample from the mid- or later flow of urine in a sterile container.
(v) Remove catheter upon completion of the procedure.
Indicate on the test request that this is a catheterised specimen.
(i) Clean the catheter collection port with a 70% alcohol wipe.
(ii) Using aseptic technique, puncture the collection port with a needle attached to a syringe. (Do not collect urine from collection bag).
(iii) Aspirate the urine, and place it in a sterile container.
SPA is useful in determining urinary infection in adults in whom infection is suspected, and for whom results from routine procedures have been equivocal and diagnosis is critical. SPA is also useful in paediatric patients when clean-catch urine samples are difficult to obtain.
(i) Shave if necessary, and disinfect the suprapubic skin overlying the urinary bladder.
(ii) Make a lance wound through the epidermis at the midline just above the symphysis pubis.
(iii) Introduce the needle and aspirate urine from the bladder and place specimen in a sterile container.
The bladder washout test is useful in determining the site of infection in the urinary tract. Results are equivocal in about 10 to 20% of patients.
(i) Clean the urethral area with soapy water and rinse the area well with water.
(ii) Insert an indwelling catheter into the bladder through the urethra.
(iii) Collect an initial urine sample into a sterile container and refrigerate it.
(iv) Empty the bladder through the urethral catheter and then irrigate it using sterile non-bacteriostatic 0.85% NaCl.
(v) Collect three additional samples (5 – 10 mL each) at 10 minutes interval into separately labelled containers after irrigation of the bladder is performed.
(vi) Submit the initial and timed collection samples to the laboratory, clearly labelled with the time of each specimen collection.
This is useful for determining the site of infection in the urinary tract.
(i) The patient is to have a full bladder before performing cystoscopy.
(ii) Clean the urethral area with soapy water and rinse the area well with water.
(iii) Insert a cystoscope (obturator in place) into the bladder.
(iv) With aseptic technique, collect 5 – 10 mL of urine from open stopcock into a sterile container.
(v) Label this sample CB (catheterised bladder urine) and refrigerate it. Then irrigate the bladder using sterile non-bacteriostatic 0.85% NaCl.
(vi) After irrigation of the bladder and insertion of the ureteral catheters, collect irrigating fluid passing from the bladder through the ureteral catheters by holding the ends of both catheters over an opened sterile container.
(vii) Label this WB (washed bladder urine) and refrigerate it.
(viii) Pass the ureteral catheters to each midureter or renal pelvis without introducing additional irrigating fluid. Open both stopcocks of the cystoscope to empty the bladder.
(ix) Discard the first 5 – 10 mL of urine from each ureteral catheter.
(x) Collect four consecutive paired urine specimens (5 – 10 mL) directly into opened sterile containers.
(xi) Label these samples LK-1, RK-1, LK-2, RK-2 (LK for left kidney and RK for right kidney). Submit all samples to the laboratory for culture.
Indicated in the diagnosis of chronic prostatitis, but contraindicated in the setting of acute prostatitis (as there is a risk of bacteraemia).
(i) Patient should have a full bladder before performing the procedure.
(ii) Retract foreskin (if necessary) and cleanse penis as described in Clean Catch Urine Collection (male).
(iii) Collect pre-massage midstream Urine Specimen into a sterile container.
(iv) Once patient stops voiding, insert gloved finger per rectum with lubrication.
(v) Massage prostate from periphery to midline.
(vi) Collect a few drops of secretion from urethra into a second sterile container.
(vii) Collect 10 mL of urine for post-massage Urine Culture into a third sterile container.
(viii) Label these samples appropriately and submit them to the laboratory for culture.
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