3.4 Guidance for collecting and transporting specific samples

Only a proportion of blood cultures yield significant microorganisms. The proportion will depend on who is sampled and local infection prevalence, but usually, fewer than 10% of samples will grow significant pathogens.

Even with good collection technique, 1–3% of blood cultures are contaminated.

Blood culture contamination rates are minimised by strict adherence to aseptic collection technique and, whenever possible, collecting peripheral blood via venepuncture with proper skin antiseptic preparation, for example with 70% alcohol, 0.5% chlorhexidine gluconates or 10% povidone-iodine.

The total volume of blood for blood culture request is the most important aspect, because bloodstream infections have a low concentration of organisms in the blood, estimated as 99% of pathogens, but in practice it is very difficult to obtain this volume.

For optimal sampling in adults, it is recommended that 20–30 ml of blood is taken per set (a set means an aerobic bottle plus an anaerobic bottle), and that two sets (four bottles in total) are taken. While this is ideal, local decisions may need to be made based on sampling costs.

For suspected infective endocarditis, three sets of blood cultures are recommended, taken from different sites one to two hours apart. This is because endocarditis may be caused by normal skin flora (especially prosthetic valve endocarditis) and because patients will need to be on a long course of antibiotics. It is therefore vital to make sure that the diagnosis and causative organism are correctly identified: the results of three or four positive blood cultures with the same organism establishes the presence of continuous bacteraemia and helps the physician determine the clinical relevance of the isolates.

In children, the optimal volume of blood is less well-prescribed; and especially for infants, the need for volume to improve test sensitivity needs to be balanced with the risk of over-bleeding the infant. The recommended volume is therefore based on the weight of the child: for infants less than 1–2 kg, 2 ml should be submitted; for a child of 2–12 kg, 4 ml; and more than 12 kg, 10 ml. A single aerobic bottle should be used unless anaerobic infection is suspected. Alternatively, a single special paediatric culture bottle can be used.

If a vascular line infection is suspected, cultures should be taken from both the line and peripherally, to determine whether the line is contaminated or infected, and whether the infection has become systemic.

Specimens should reach the laboratory with no delay. Although commercial blood culture bottles have been carefully formulated to optimise bacterial growth, prolonged transportation may result in death of fastidious bacteria and overgrowth of other bacteria. If there is going to be a long delay in sending specimens to the laboratory, the bottles should be kept in an incubator at 35–37°C in an onsite laboratory or in a safe place on the ward.

Cerebrospinal fluid (CSF) must be collected using strict aseptic technique, both to minimise specimen contamination and to prevent bacteria being introduced into the central nervous system. Either povidone-iodine or chlorhexidine can be used for disinfection.

The risk of contamination is higher when CSF is collected from catheters or shunts. Such contamination is problematic, because organisms (such as coagulase-negative Staphylococci) are likely to cause many CSF catheter and/or shunt infections: it may therefore be necessary to take several samples to help to distinguish contamination from true infection.

The volume of CSF depends on the pathogens sought. For routine bacterial cultures, 2–3 ml is adequate for adults. Note that for fungal cultures, microbial yield is more proportional than bacterial yield to the volume of cerebrospinal fluid cultured.

The laboratory should consider using sequential testing to reduce the number of unnecessary CSF tests, for example by use of a testing algorithm that specifies a chronological order that tests should be performed in. For example, the first step might be recording opening pressure to assess if inflammation is present, followed by assessment of colour, or Gram staining for bacterial meningitis. (The ddxof website provides an example of an algorithm for analysing CSF.)

When testing CSF for tuberculosis (TB) culture, samples can be sent for acid-fast smear with the important caveat that a single sample (approximately 20–40%) has low sensitivity. Several large volume (10–15 ml) lumbar punctures are often needed for a microbiological diagnosis; sensitivity increases to >85% when four spinal taps are performed. While culture can take several weeks and has low sensitivity (~40–80%), it should be performed to determine drug susceptibility (Marx and Chan, 2011).

CSF specimens should be transported immediately to the laboratory. For suspected bacterial meningitis, the laboratory should report the results of initial tests (including microscopy with cell counts and Gram stain, and latex agglutination for bacterial organism such as N. meningitidis) within 30 minutes of receipt of the specimen so that appropriate treatment can be given.

From collection through processing, CSF specimens (except aliquots collected for viral cultures, performed only in specialised centres) should not be refrigerated until initial processing is completed.

For diagnosing lower respiratory tract infections (LRTIs), expectorated sputum is the most commonly received sample, as it can be obtained easily and non-invasively. However, patients with pneumonia often have difficulty in producing a good-quality sputum sample due to pain and breathlessness, and specimens are frequently contaminated by normal resident bacteria of the oropharynx, preventing the determination of the true pathogen and leading to wrong results.

More invasive samples (such as biopsies, brushings and lavage specimens) have better sensitivity and specificity, but require bronchoscopy, which is rarely required or performed for uncomplicated pneumonia.

The value of sputum microscopy and culture in the diagnosis, management and outcome of LRTIs is therefore a matter of controversy, and many laboratories no longer process sputum samples routinely.

The quality of an expectorated sputum sample can be assessed by Gram staining to look for epithelial cells (indicative of a poor quality sample), white blood cells and organisms (indicative of infection). The average number of the different cell types in 20–30 low power fields is calculated and the total score is based on Bartlett criteria. A final score of 0 or less indicates lack of active inflammation or contamination (non-acceptable sample), and a score of 1 and above is considered an acceptable sample. Only samples with a score of 1 or above should be cultured.

A sputum Gram stain examined according to the correct guidelines is considered useful in the initial evaluation of patients with pneumonia (Del Rio-Pertuz et al., 2019). However, it is labour-intensive, and local guidelines should be developed to optimise the cost-effectiveness of processing sputum samples. An additional factor to consider is the local prevalence of TB, as there may be a risk of transmission to laboratory staff from sputum samples and appropriate precautions should be taken.

Most respiratory tract specimens are likely to contain at least a few contaminating microorganisms, so specimens should be transported rapidly to the laboratory to minimise contaminant growth. If transportation or processing is delayed for more than two hours, specimens should be rejected.

The laboratory diagnosis of enteric infections is challenging, due to the diversity of the normal flora. Local policy should decide which organisms should be tested for.

GLASS (which will be introduced later in this module) looks for Salmonella spp. and Shigella spp. due to the emergence of multidrug-resistant strains, but many laboratories will also look for Campylobacter spp. Stool specimens collected from patients who develop diarrhoea in the hospital should be tested for the presence of C. difficile toxin.

There is little value in routinely testing multiple stool specimens as part of an evaluation of acute diarrhoea, as most pathogens are detected in the first specimen. Repeat specimens should be sent if symptoms persist.

Samples should be transported to the laboratory within two hours. Shigella spp. are very sensitive, so prompt delivery to the laboratory is critical if they are to be cultured successfully (WHO, 2016).

Because urine is easily contaminated with commensal flora, careful collection of specimens is vital to minimise this. Midstream urine is ideal; first morning urine is best because it may contain more pathogenic organisms.

Patients need to be instructed on how to clean themselves and take the midstream urine. Samples from catheter tips and bags are not appropriate (Miller et al., 2018) because they are frequently colonised and/or contaminated.

The volume of urine specimens is not as important as collecting urine in sterile specimen containers, and obtaining a ‘clean catch’ or midstream urine.

Urine samples should be collected and reach the laboratory as soon as possible, and culture should commence immediately to avoid overgrowth of other non-pathogenic organisms. Specimens that won’t reach the laboratory within two hours of collection can be refrigerated for up to 24 hours.

Preservatives such as boric acid, which inhibits bacterial growth and prevents overgrowth with contaminants, may be used for storage and transportation of urine. Non-buffered boric acid is preferred so as to reduce the effect of the preservative on the actual pathogens (Weinstein, 1983).