Free ASCP MLS Exam Practice Questions Mock Test: Part 15 – Microbiology Preanalytic Procedures

• Best practice: Use a sterile syringe to puncture the catheter’s sampling port (after disinfecting with alcohol/iodine).

• Why?

  • Avoids contamination from the drainage bag (older urine, colonized bacteria).

  • Prevents disruption of the closed catheter system (reduces infection risk).

Why Not Other Methods?

• a) Incorrect – Cutting the catheter tip contaminates the specimen and damages the device.

• b) Incorrect – Disconnecting the catheter introduces contamination and breaches infection control.

• c) Incorrect – Drainage bag urine is stagnant and unreliable for culture (high false-positive rate).

Preanalytic variables are factors that affect test results before the specimen is analyzed in the lab. These occur during collection, handling, transport, and storage.

Why Option (c) is NOT a Preanalytic Variable:

• Duration of incubation is part of the analytic phase (laboratory processing).

• Once the specimen reaches the lab, incubation time is controlled by lab protocols, not by pre-collection or transport factors.

Why the Other Options ARE Preanalytic Variables:

• a) Site selection for specimen collection (e.g., avoiding contaminated areas) affects specimen quality.

• b) Method of specimen transport (e.g., use of preservatives, temperature control) impacts microbe survival.

• d) Timing of specimen collection (e.g., during fever for blood cultures, early morning sputum) influences pathogen detection.

The preanalytic phase (before testing begins) is the most error-prone stage in microbiology, and specimen integrity is critical. Key factors include:

1. Proper collection technique (e.g., sterile site avoidance of contamination).

2. Timely transport (prevents overgrowth of commensals or pathogen death).

3. Appropriate storage (e.g., anaerobic specimens must avoid oxygen).

Errors here lead to:

• False negatives (e.g., pathogens die due to delay).

• False positives (e.g., contamination with normal flora).

Why Not the Other Options?

• a) Correct antibiotic therapy → Post-analytic (treatment follows results).

• c) Selection of culture media → Analytic phase (lab’s responsibility).

• d) Incubation temperature → Analytic phase (controlled by the lab).

False-negative stool cultures often result from:

1. Prior antibiotic use:

   • Antibiotics suppress growth of Salmonella, Shigella, Campylobacter, and other enteric pathogens.

   • Even partial treatment can reduce bacterial load below detectable levels.

Why Not the Other Options?

• a) Refrigerating promptly → Preserves pathogens (recommended if delay >1 hour).

• b) Cary-Blair medium → Optimal for transport (maintains viability of enteric bacteria).

• d) Sterile container → Standard practice (avoids contamination, doesn’t cause false negatives).

Transport media serve a critical role in microbiology by maintaining the specimen in a state that:

• Preserves pathogen viability (keeps microorganisms alive).

• Prevents overgrowth (does not encourage rapid multiplication).

Why Option (c) is Correct:

• Many clinical specimens (e.g., throat swabs, genital samples, tissue biopsies) cannot be processed immediately.

• Transport media (e.g., Amies, Stuart, Cary-Blair) contain buffers and nutrients to:

  • Prevent drying (which kills delicate bacteria like Neisseria gonorrhoeae).

  • Maintain pH and osmotic balance (preventing cell lysis).

  • Inhibit overgrowth of commensals (e.g., throat flora masking Corynebacterium diphtheriae).

Why Other Options Are Incorrect:

• a) To sterilize the specimen → Incorrect, because sterilization kills all microbes (transport media aim to preserve them).

• b) To promote rapid bacterial growth → Incorrect, as transport media are designed to maintain but not amplify bacterial numbers (culture media are used for growth).

• d) To eliminate contaminants → Incorrect, because while some selective transport media (e.g., for Bordetella pertussis) suppress contaminants, most general-purpose media do not.

• Sodium polyanetholsulfonate (SPS) is an anticoagulant and antimicrobial inhibitor added to blood culture bottles.

• Key functions:

  1. Neutralizes host defenses:

     • Inactivates complement (reduces bacterial lysis).

     • Inhibits phagocytosis by neutrophils.

  2. Anticoagulant: Prevents clotting (ensures bacteria remain free in broth).

  3. Counteracts antibiotics: Mild effect (but primarily β-mercaptoethanol is used for this).

Why Not Other Options?

• a) Incorrect – SPS has weak antibiotic-neutralizing effects (other additives like resin/charcoal are better).

• b) Incorrect – SPS prevents clotting, not RBC clumping (which is addressed by EDTA).

• d) Incorrect – SPS does not specifically aid anaerobes (but anaerobic bottles support their growth).

Contamination in blood cultures most often occurs due to inadequate skin disinfection, which allows commensal skin flora (e.g., Staphylococcus epidermidis, Cutibacterium acnes) to enter the culture bottle. Key factors:

1. Incomplete disinfection:

   • Failure to use alcohol + iodine/chlorhexidine sequentially.

   • Insufficient contact time (e.g., <30 seconds for alcohol).

2. Touch contamination:

   • Re-palpating the venipuncture site after disinfection.

Why Not the Other Options?

• a) Non-sterile needles → Rare (modern needles are pre-sterilized).

• c) Inadequate blood volume → Reduces sensitivity but doesn’t cause contamination.

• d) Delayed transport → May reduce viability but doesn’t introduce contaminants.

• Optimal anaerobic wound culture requires:

  1. Pus or tissue (not swabs) – Swabs expose anaerobes to oxygen, killing them.

  2. Syringe aspiration – Maintains an oxygen-free environment (anaerobic transport).

  3. Before antibiotics – Antibiotics reduce bacterial viability, causing false negatives.

Why Not Other Options?

• a) Swabs before antibiotics – Swabs are suboptimal (air exposure kills anaerobes).

• b) Swabs after antibiotics – Antibiotics + swab = worst yield.

• d) Syringe after antibiotics – Antibiotics reduce recovery, even with proper collection.

For Clostridioides difficile toxin testing, proper handling is critical to prevent toxin degradation:

1. Refrigeration (2–8°C):

   • Preserves toxin stability if testing is delayed (>2 hours).

   • Prevents overgrowth of commensal bacteria that may degrade toxins.

2. Avoid:

   • Room temperature storage → Toxins degrade rapidly (false negatives).

   • Freezing → May lyse cells and inactivate toxins.

Why Not the Other Options?

• a) Preservative → Not recommended (may interfere with PCR/ELISA tests).

• c) Saline solution → Dilutes toxins (invalidates results).

• d) Room temperature for 24h → Toxin degradation guaranteed.

Anaerobic bacteria die in the presence of oxygen, so tissue biopsies must be transported in:

1. Anaerobic transport vials:

   • Pre-reduced, oxygen-free environment (often containing anaerobic gas mix).

   • Maintains viability of fastidious anaerobes (e.g., Bacteroides, Clostridium).

2. Alternatives:

   • Sterile syringe (if immediately injected into anaerobic broth).

Why Not the Other Options?

• a) Dry sterile swab → Oxygen exposure kills anaerobes (swabs trap air).

• c) Formalin-filled container → Preserves tissue for pathology but kills bacteria.

• d) Non-sterile plastic bag → Contamination risk + oxygen exposure.

Urine specimens for culture must be processed within 2 hours of collection if unpreserved, or within 24 hours if refrigerated (4°C) or preserved with boric acid.

• >24 hours at room temperature:

  • Overgrowth of contaminants (e.g., Lactobacillus, skin flora) occurs, leading to false-positive results.

  • Pathogens may die (e.g., Neisseria gonorrhoeae, Streptococcus pneumoniae), causing false negatives.

Why Not the Other Options?

• a) Catheterization → Acceptable method (avoids vaginal/perianal contamination).

• c) Clean-catch midstream → Gold standard for reducing contamination.

• d) Boric acid preservation → Extends viability to 24 hours (not a rejection reason).

In suspected septicemia, bacteria are more likely to be present in the bloodstream during fever spikes (due to increased bacterial load). Collecting blood cultures at this time improves the chances of detecting pathogens, maximizing diagnostic yield.

Why the other options are incorrect:

• a) Incorrect – Hemolysis is unrelated to timing.

• c) Incorrect – The number of bottles depends on protocol, not timing.

• d) Incorrect – Contaminants are minimized via proper skin antisepsis, not timing.

• Legionella pneumophila requires BCYE agar for growth because:

  • Charcoal absorbs toxic metabolites.

  • Yeast extract provides nutrients (L-cysteine, iron).

  • Antibiotics (e.g., cefamandole, polymyxin) suppress normal flora.

Why Not Other Options?

• a) Thiosulfate citrate bile salts (TCBS) – Used for Vibrio, not Legionella.

• b) Anaerobic incubation – Legionella is aerobic/facultative.

• c) Rejecting the specimen – Bronchoscopy samples are superior to sputum for Legionella (lower contamination risk).

Delayed transport of stool specimens for ova and parasite (O&P) examination can compromise results in two key ways:

1. Death and distortion of trophozoites:

   • Trophozoites (motile, fragile forms of protozoa like Giardia or Entamoeba) rapidly degrade outside the body, making them difficult to identify microscopically.

   • Cysts (hardier forms) survive longer but may also distort over time.

2. Increased bacterial overgrowth:

   • Bacterial proliferation in unpreserved stool can:

     • Obfuscate parasites under microscopy.

     • Alter stool pH, accelerating parasite disintegration.

Why Not the Other Options?

• a) Increased parasite motility → False; motility decreases with time due to cooling/oxygen exposure.

• b) and c) → Partially correct, but d) covers both critical issues.

• Squamous epithelial cells (>25/low-power field) indicate saliva contamination, meaning the specimen is not true sputum but mostly oropharyngeal flora.

• Mixed bacteria (e.g., Streptococcus viridans, Neisseria) further suggest normal mouth flora, not a lower respiratory infection.

Why Not Other Options?

• a) Pneumococcal pneumonia – Expect PMNs + gram-positive diplococci, not squamous cells.

• b) Anaerobic infection – Requires sterile-site specimens (e.g., abscess fluid), not saliva.

• c) Haemophilus pneumonia – Expect PMNs + gram-negative coccobacilli, not mixed flora.

Cary-Blair medium is the gold-standard transport medium for stool samples suspected of containing Vibrio cholerae (and other enteric pathogens like Salmonella, Shigella, Campylobacter) because:

1. Preserves viability:

   • Alkaline pH (~8.4) and reducing agents maintain Vibrio survival.

2. Prevents overgrowth:

   • Inhibits commensal bacteria while preserving pathogens.

3. WHO-recommended:

   • Specifically endorsed for cholera outbreak investigations.

Why Not the Other Options?

• b) Stuart’s medium → For genital/nasopharyngeal swabs (not stool).

• c) Amies medium → For aerobic bacteria/swabs (e.g., Neisseria), not Vibrio.

• d) Buffered glycerol saline → Outdated; inhibits Vibrio and degrades quickly.

False-positive blood cultures (growth of non-pathogenic bacteria) most often result from:

1. Skin flora contamination:

   • Commensals like Staphylococcus epidermidis, Cutibacterium acnes, or Corynebacterium spp. are introduced during venipuncture.

   • Causes:

     • Inadequate skin disinfection (e.g., skipped iodine/chlorhexidine step).

     • Re-palpating the site after disinfection.

Why Not the Other Options?

• a) Low blood volume → Causes false negatives (insufficient pathogen load), not false positives.

• b) Povidone-iodine → Reduces contamination (when used correctly).

• d) Delay in incubation → May cause false negatives (pathogen death), not false positives.

• The ideal dilution for blood cultures is 1:5 to 1:10 (e.g., 8–10 mL blood in 80–100 mL broth).

• This ratio:

  • Dilutes natural inhibitors (e.g., antibodies, complement, antimicrobial drugs).

  • Prevents false negatives from bacterial overgrowth suppression.

  • Optimizes nutrient availability for microbial growth.

Why Not Other Ratios?

• a) 1:2 – Too concentrated; inhibitors may remain active.

• b) 1:3 – Still insufficient dilution for optimal recovery.

• d) 1:30 – Over-dilution may reduce bacterial detection sensitivity.

Anaerobic cultures require specimens from normally sterile sites where anaerobes thrive (e.g., abscesses, deep tissues). Vaginal swabs are unsuitable because:

1. Heavily contaminated with normal flora:

   • Vaginal microbiota include facultative anaerobes (e.g., Lactobacillus) and aerobes, which overgrow true anaerobes.

2. Oxygen exposure:

   • Swabs trap air, killing obligate anaerobes (e.g., Bacteroides, Prevotella).

Why the Other Options Are Suitable:

• b) Abscess aspirate → Ideal (anaerobes thrive in pus).

• c) Deep tissue biopsy → Protected from oxygen.

• d) Anaerobic blood culture bottle → Designed for bloodborne anaerobes.

Boric acid is the preferred preservative for urine cultures when delays in processing are expected (e.g., during transport or storage). Here’s why:

1. Maintains bacterial viability:

   • Inhibits overgrowth of contaminants while preserving pathogenic bacteria (e.g., E. coli, Enterococcus).

2. Stabilizes specimen:

   • Prevents pH shifts and bacterial multiplication for up to 24 hours at room temperature.

3. Clinically validated:

   • Recommended by CLSI guidelines for urine culture preservation.

Why Not the Other Options?

• b) Formalin → Kills bacteria (used for cytology/parasitology, not cultures).

• c) EDTA → Anticoagulant (used for blood/CSF, not urine cultures).

• d) Sodium fluoride → Inhibits glycolysis (used for glucose testing, not cultures).

• TCBS agar is selective for Vibrio species (e.g., V. cholerae, V. parahaemolyticus).

• Key features:

  • High pH (8.6–9.0) inhibits most gut flora.

  • Sucrose fermentation helps differentiate Vibrio (yellow colonies = sucrose fermenters like V. cholerae; green = non-fermenters like V. parahaemolyticus).

Why Not Other Options?

• a) Colistin-nalidixic acid (CNA) – Used for Listeria (not stool pathogens).

• b) MacConkey-sorbitol – Used for E. coli O157:H7 (sorbitol non-fermenter), not Campylobacter.

• c) Mannitol salt agar – Selective for Staphylococcus (not enteric pathogens).

Proper specimen labeling is critical in clinical microbiology to ensure accurate diagnosis, patient safety, and reliable test results. A specimen labeled only as “body fluid” is insufficient because:

1. Lack of Patient Identification: It does not include essential details such as the patient’s name, medical record number, or date of birth, which could lead to misidentification.

2. Unknown Source: Different body fluids (e.g., pleural fluid, synovial fluid, cerebrospinal fluid) require specific processing protocols and tests. Without knowing the source, the lab cannot perform appropriate testing.

3. Quality and Safety Concerns: Improperly labeled specimens increase the risk of errors, delayed diagnosis, and potential harm to the patient.

Why Other Options Are Incorrect:

• b) Processed immediately – Processing an improperly labeled specimen could lead to incorrect testing or reporting.

• c) Sent for routine culture only – Without knowing the source, routine culture may be inappropriate (e.g., CSF requires specialized testing).

• d) Stored until source is identified – Storing an unlabeled specimen is not a solution; the specimen should be rejected, and a properly labeled one should be requested.

• Neutrophil nuclei appear dark blue/purple in Gram stain when:

  • Decolorization (alcohol/acetone step) is too short – Excess crystal violet remains, overstaining cells.

  • Overuse of crystal violet – Can also cause excessive dye retention.

Why Not Other Options?

• a) Iodine omitted – Would cause all bacteria to stain pink (Gram’s iodine fixes crystal violet; without it, dye washes out).

• c) Smear too thin – Leads to faint staining, not dark nuclei.

• d) Normal staining – Neutrophil nuclei should be pale pink/red (counterstained by safranin).

Respiratory specimens (e.g., sputum, bronchoalveolar lavage) must be processed within 2 hours of collection if stored at room temperature because:

1. Fastidious pathogens die:

   • Bacteria like Streptococcus pneumoniae and Haemophilus influenzae degrade rapidly.

2. Overgrowth of commensals:

   • Oral flora (e.g., Streptococcus viridans) can multiply, obscuring true pathogens.

Exceptions:

• Refrigeration (4°C) extends viability to 24 hours (but some pathogens like H. influenzae still decline).

• Preserved specimens (e.g., in transport media) may tolerate slightly longer delays.

Why Not the Other Options?

• a) 1 hour → Too strict (most labs allow up to 2 hours).

• c) 6 hours → Unacceptable (significant pathogen loss).

• d) 24 hours → Only valid for refrigerated (not RT) specimens.

False-positive fecal occult blood tests (FOBT) can occur due to:

1. Dietary peroxidases:

   • Red meat (especially rare/undercooked) contains hemoglobin/myoglobin, which reacts with guaiac-based FOBT (gFOBT).

   • Certain raw vegetables (e.g., horseradish, radishes) also contain peroxidases.

2. Other causes:

   • NSAIDs/aspirin (cause GI bleeding).

   • Menstrual blood contamination.

Why Not the Other Options?

• a) 2-hour delay → Minimal impact (false negatives are more likely with delays).

• b) High vitamin C → Causes false negatives (inhibits chemical reaction in gFOBT).

• d) Cary-Blair medium → Used for bacterial stool cultures, not FOBT.

Collecting multiple sets of blood cultures (typically 2–4 sets) is critical because:

1. Intermittent bacteremia:

   • Pathogens may not be continuously present in blood (e.g., early sepsis, endocarditis).

   • Multiple sets improve detection (sensitivity rises from ~80% to >95%).

2. Volume matters:

   • Each set adds blood volume, increasing the chance of capturing low-level bacteremia.

Why Not the Other Options?

• a) Detect contamination → Helps but is secondary (skin flora are usually sporadic).

• b) Identify resistance → Requires culture growth first, not collection strategy.

• d) Reduce costs → False; multiple sets increase costs but are clinically justified.

For closed abscesses, needle aspiration is the gold standard because:

1. Minimizes contamination:

   • Directly samples purulent material without exposing it to skin flora.

2. Preserves anaerobes:

   • Syringe can be sealed to exclude oxygen (critical for anaerobes like Bacteroides).

3. Higher diagnostic yield:

   • Avoids dilution or interference from surface contaminants.

Why Not the Other Options?

• a) Swab after incision → Contaminates specimen with skin flora.

• c) Cotton swab from tissue → Superficial/inadequate for deep abscesses.

• d) Gauze drainage → Exposed to air/skin, useless for culture.

• Urine is an acceptable specimen for AFB culture when renal/urogenital tuberculosis is suspected (Mycobacterium tuberculosis can be detected in urine).

• First-morning, midstream urine (3 consecutive days) is ideal for maximizing yield.

Why Not Other Options?

• a) Feces for anaerobic culture – Unacceptable (normal gut flora includes anaerobes; no diagnostic value).

• b) Foley tip for aerobic culture – Unacceptable (catheter tips are not reliable for UTI diagnosis; use fresh urine instead).

• c) Rectal swab for gonococci smear – Unacceptable (Neisseria gonorrhoeae requires mucosal swabs (urethral/cervical), not stool).

Cerebrospinal fluid (CSF) is a critical and sterile body fluid, especially when evaluating for conditions like meningitis. Delaying processing can lead to:

• Loss of viability of fastidious organisms (e.g., Neisseria meningitidis)

• Overgrowth of contaminants

• Decreased sensitivity of culture and Gram stain

Why the other options are less appropriate:

• a) 15 minutes 
  While 15 minutes is a short delay, even this should be avoided when possible for optimal organism recovery.

• b) 1 hour 
  Delaying up to 1 hour can compromise the integrity of sensitive organisms.

• c) 4 hours refrigerated

For fungal bloodstream infections (e.g., Candida, Cryptococcus), the lysis-centrifugation method (Isolator™ tube) significantly improves recovery because:

1. Mechanism:

   • Lyses blood cells and concentrates fungi via centrifugation.

   • Directly plates sediment on fungal media (e.g., Sabouraud agar).

2. Advantages:

   • Higher sensitivity than conventional blood cultures (especially for Histoplasma, Malassezia).

   • Faster detection (fungi grow on solid media sooner).

Why Not the Other Options?

• a) Smaller blood volume → Reduces sensitivity (fungemia often low-grade).

• b) Mycobacterial broth → For TB, not fungi.

• d) Anaerobic media only → Fungi are aerobic/facultative (require aerobic conditions).

• Stool specimens for enteric pathogens (e.g., Salmonella, Shigella, Campylobacter) should ideally be processed immediately.

• If processing is delayed beyond 1–2 hours, refrigeration (2–8°C) helps preserve bacterial viability while preventing overgrowth of normal flora.

Why not other options?

• a) Frozen immediately – Freezing can damage fragile pathogens (e.g., Campylobacter).

• c) Incubated at 37°C – Promotes rapid overgrowth of commensals, masking pathogens.

• d) Room temperature indefinitely – Leads to bacterial degradation and false negatives.

Wound specimens may be rejected if:

• Insufficient clinical details (e.g., no wound type/site) hinder proper interpretation.

• Dry swabs can lead to bacterial desiccation and false-negative results.

Other options:

• a) Correct collection method (sterile needle/syringe is ideal).

• c) Rapid transport (30 minutes) is acceptable.

• d) Anaerobic transport medium is appropriate if anaerobes are suspected.

Anaerobic cultures require specimens protected from oxygen exposure, as anaerobes die rapidly in air. The least acceptable specimen is:

• Swab from a superficial wound exposed to air:

  • Swabs trap oxygen, killing anaerobes.

  • Superficial wounds are likely contaminated with aerobes, masking anaerobic pathogens.

Why the Other Options Are Better Choices:

• a) Aspirated pus from a sterile site → Ideal (no oxygen exposure, high yield).

• b) Tissue biopsy in oxygen-free vial → Excellent (maintains anaerobe viability).

• d) Synovial fluid by needle aspiration → Acceptable (if transported anaerobically).

Anticoagulants (e.g., EDTA, citrate, heparin) are avoided in blood cultures because:

1. Bacterial growth inhibition:

   • EDTA chelates magnesium and calcium, which are essential for bacterial metabolism.

   • SPS (sodium polyanethol sulfonate), the preferred anticoagulant in blood culture bottles, is less inhibitory.

2. False-negative risk:

   • Fastidious organisms (e.g., Neisseria meningitidis, Streptococcus pneumoniae) may fail to grow.

Why Not the Other Options?

• b) Promote clot formation → False; anticoagulants prevent clotting.

• c) Interfere with Gram staining → Rare; SPS can inhibit PCR but not Gram stains.

• d) Increase hemolysis → Unrelated; hemolysis depends on collection technique, not anticoagulants.

• Male urethral discharge is typically cultured for sexually transmitted pathogens, primarily:

  • Neisseria gonorrhoeae (fastidious, requires Thayer-Martin agar with antibiotics).

  • Other fastidious organisms (e.g., Haemophilus), which grow on chocolate agar (heated blood for V/X factors).

Why Not Other Options?

• a) Blood + phenylethyl alcohol (PEA) – Used for anaerobes/gram-positives (not STIs).

• b) EMB + blood – Routine for enterics (e.g., E. coli), not gonorrhea.

• c) Thioglycolate broth – Enhances anaerobe recovery (irrelevant here).

To minimize contamination during urine culture collection, the clean-catch midstream technique is essential because:

1. Discards the initial stream:

   • The first portion of urine washes away urethral commensals (e.g., Lactobacillus, Staphylococcus epidermidis).

2. Collects midstream urine:

   • Represents bladder urine with minimal contamination.

3. Sterile container:

   • Prevents introduction of environmental bacteria.

Why Not the Other Options?

• a) First portion → Contaminated with urethral flora.

• c) Room temperature storage → Promotes bacterial overgrowth (false positives).

• d) Non-sterile container → Introduces contaminants.

Anaerobic bacteria die in the presence of oxygen, so specimens must be collected and transported to minimize air exposure:

1. Aspirates (e.g., abscess fluid, tissue):

   • Collected via needle/syringe to exclude air.

   • Immediately transferred to anaerobic transport media (or injected into blood culture bottles).

2. Avoid swabs/open-air collection:

   • Swabs (even in transport media) often retain oxygen, reducing anaerobic recovery.

Why Not the Other Options?

• a) Swab specimen in open air → Oxygen exposure kills anaerobes (e.g., Bacteroides, Clostridium).

• c) Urine in clean catch container → Irrelevant for anaerobes (urine cultures target aerobes like E. coli).

• d) Saliva in sterile cup → Contaminated with oral aerobes; anaerobes are undetectable this way.

• Ziehl-Neelsen (ZN) stain uses heat (steam) to drive carbol fuchsin into the waxy mycolic acid layers of acid-fast bacteria (e.g., Mycobacterium tuberculosis).

• Kinyoun stain is “cold” – It uses a higher concentration of phenol in carbol fuchsin to penetrate without heating.

Why Not Other Options?

• a) Dye type – Both use carbol fuchsin (same dye).

• b) Microscope used – Both are viewed under brightfield microscopy.

• c) Acid decolorizer strength – Both use 3% HCl in ethanol (same decolorizer).

• Parasites (e.g., Giardia, Entamoeba, worms) are not consistently shed in every stool sample.

• Intermittent shedding means a single specimen has a high false-negative rate.

• Collecting 3 specimens over separate days (or every other day) increases detection sensitivity by ~90% (vs. ~50% for one sample).

Why not other options?

• b) Incorrect – Multiple collections increase costs, but this is justified for accuracy.

• c) Incorrect – Bacterial overgrowth is irrelevant; parasites are detected via microscopy/antigen tests.

• d) Incorrect – Volume matters less than timing (shedding cycles).

For suspected diphtheria (caused by Corynebacterium diphtheriae), proper specimen collection and handling are crucial for accurate diagnosis:

Why Option (b) is Correct:

1. Sterile Swab:

   • A sterile swab (preferably Dacron or rayon) should be used to avoid contamination and ensure optimal recovery of C. diphtheriae.

   • Cotton swabs may contain fatty acids that inhibit bacterial growth.

2. Immediate Transport:

   • C. diphtheriae is fastidious and sensitive to drying.

   • The swab should be transported quickly to the lab (within 2 hours) to maintain viability.

   • If delay is unavoidable, use transport media (e.g., Amies or Stuart medium).

Why Other Options Are Incorrect:

• a) Collected after initiation of antibiotics → Antibiotics can suppress bacterial growth, leading to false-negative results.

• c) Stored at room temperature for 12 hours → Prolonged storage without proper transport media reduces bacterial viability.

• d) Collected in a dry container without transport medium → C. diphtheriae may die due to desiccation; transport media is preferred.

The optimal blood-to-broth ratio in blood culture bottles is 1:5 to 1:10 (e.g., 8–10 mL blood in 50–100 mL broth). This balance is critical because:

1. Dilution effect:

   • Too little blood (e.g., 1:100) → Dilutes pathogens below detectable levels.

   • Too much blood (e.g., 1:1) → Overwhelms the broth’s anticoagulant (SPS) and nutrients.

2. Neutralizes inhibitors:

   • Human serum contains antibodies/complement that kill bacteria; dilution minimizes this.

Why Not the Other Options?

• a) 1:1 → Excessive blood (causes clotting, false negatives).

• b) 1:5 → Acceptable but less common (most commercial bottles use ~1:10).

• d) 1:100 → Pathogens undetectable due to over-dilution.

The gold-standard antiseptic protocol for blood culture collection involves two-step disinfection to minimize contamination:

1. Povidone-iodine (10%):

   • Broad-spectrum antimicrobial activity (kills bacteria, fungi, and spores).

   • Applied in a circular motion for 30–60 seconds, then allowed to dry.

2. 70% isopropyl alcohol:

   • Removes iodine residue (which can interfere with some culture systems) and provides rapid bacterial kill.

Why Not the Other Options?

• a) 70% alcohol only → Less effective against spores and fungi (higher contamination risk).

• c) Soap and water → Cleans but does not sterilize the skin.

• d) Hydrogen peroxide → Not recommended for blood cultures (poor efficacy, tissue irritation).

Key Alternatives:

• Chlorhexidine (2%) + alcohol is equally effective (and preferred in iodine-allergic patients).

• Campylobacter jejuni requires:

  1. Selective media (e.g., Campy-CVA, Skirrow’s, or CCDA agar) with antibiotics to suppress normal flora.

  2. Microaerophilic conditions (5% O₂, 10% CO₂, 85% N₂) – Mimics intestinal environment.

  3. 42°C incubation – Enhances growth while inhibiting competitors.

Why Not Other Options?

• b) Tryptic soy broth – No selectivity; delays plating reduces recovery.

• c) 35°C + room temp – C. jejuni fails to grow at these temps.

• d) Cary-Blair – A transport medium (preserves but doesn’t grow bacteria).

Viral specimens (e.g., nasopharyngeal swabs for influenza, SARS-CoV-2) require specialized transport conditions to maintain viral viability:

1. Inappropriate for viruses:

   • Dry sterile swabs → No moisture or preservatives → Rapid viral degradation (RNA/DNA breakdown).

2. Appropriate alternatives:

   • Viral transport medium (VTM) (option a/c): Contains stabilizers (e.g., protein, buffers) to preserve nucleic acids.

   • Refrigeration (2–8°C) (option d): Slows degradation if testing is delayed.

Why Not the Other Options?

• a/c) VTM/universal medium → Gold standard for viral transport.

• d) Cryovial at 2–8°C → Acceptable for short-term storage.

For accurate culture results, the catheter tip must be removed aseptically to avoid contamination with skin flora or environmental bacteria.

• a) Incorrect – Formalin preserves tissue but kills bacteria, making culture impossible.

• c) Incorrect – Delayed processing can lead to bacterial overgrowth or death, affecting results.

• d) Incorrect – A dry, sterile container is required to prevent contamination.

For pediatric blood cultures, the recommended volume balances detection sensitivity with patient safety:

1. Optimal volume:

   • Infants/neonates: 1–2 mL per bottle (total 2–4 mL per set).

   • Children (>1 month): 2–5 mL per bottle (total 4–10 mL per set).

2. Why not more?

   • Excessive volume risks anemia in small patients.

   • 1 mL of blood per kg of body weight is a general guideline (never exceed 5% of total blood volume).

Why Not the Other Options?

• a) 0.5–1 mL → Too low (misses low-level bacteremia).

• c) 5–10 mL → Excessive for most pediatric patients.

• d) 20–30 mL → Adult volumes, unsafe for children.

For adult blood cultures, the optimal volume per bottle is:

• 8–10 mL per bottle (total 16–20 mL per set, split between aerobic/anaerobic bottles).

• Why this volume?

  • <5 mL: Low sensitivity (misses bacteremia).

  • >10 mL: Risk of false positives (due to excessive anticoagulant or dilution).

Why Not the Other Options?

• a) 1–2 mL → Insufficient (yields false negatives).

• c) 20–30 mL → Excessive (may dilute the specimen or overwhelm the broth).

• d) 50 mL → Never used (cultures require ratio of blood to broth for optimal growth).

• Virus Transport Medium (VTM) must:

  1. Preserve viral viability (e.g., protein stabilizers like albumin/gelatin).

  2. Prevent bacterial/fungal overgrowth (antibiotics like gentamicin/amphotericin B).

  3. Maintain pH balance (buffered saline).

Why Not Other Options?

• a) Incorrect – VTM preserves, not grows viruses (culture requires live cells).

• c) Incorrect – VTM aims to protect all viruses, not selectively destroy any.

• d) Incorrect – Complement antibodies are irrelevant; VTM focuses on microbial inhibition.

Fastidious organisms like Neisseria gonorrhoeae require specialized transport conditions to survive until culture:

1. Amies medium with charcoal:

   • Maintains viability by preventing desiccation and toxic metabolite buildup.

   • Charcoal neutralizes inhibitory substances (e.g., fatty acids, peroxides).

   • Stuart’s or Amies are the gold standards for N. gonorrhoeae.

Why Not the Other Options?

• a) Dry cotton swab → No moisture or nutrients; pathogens die rapidly.

• c) Normal saline → No buffering/nutrients; supports only short-term survival (≤1 hour).

• d) Sterile water → Hypotonic; lyses bacterial cells.

• Blood volume is the most critical factor in detecting bacteremia.

• Adults: 20–30 mL total per culture set (2–3 bottles, 8–10 mL each).

• Why? Higher volume increases bacterial yield (each mL adds ~3% sensitivity).

Why Not Other Options?

• a) Subculturing at day 5 – Most pathogens grow within 24–48h; prolonged incubation rarely helps.

• c) Daily cultures – Useful for endocarditis, but volume per draw matters more.

• d) Multiple sets from one venipuncture – Separate venipunctures reduce contamination risk.

For lower respiratory tract infections (LRTIs), the best specimen is one that bypasses upper airway contamination and directly samples the alveoli/small airways:

1. Bronchoalveolar lavage (BAL):

   • Collected via bronchoscopy, ensuring lower airway origin.

   • Minimizes oral/nasopharyngeal contamination (unlike sputum).

   • Gold standard for pneumonia (e.g., Pneumocystis jirovecii, bacterial/fungal infections).

Why Not the Other Options?

• a) Saliva → Unacceptable (represents oral flora, not lung pathogens).

• b) Expectorated sputum → Often contaminated with upper airway flora; requires microscopic QC (e.g., <10 SECs/LPF).

• d) Nasopharyngeal swab → Detects upper respiratory pathogens (e.g., influenza, Bordetella), not LRTIs.

Sputum specimens are rejected for culture if they show excessive squamous epithelial cells (SECs), indicating saliva contamination rather than lower respiratory tract secretions. Key criteria:

• >10 SECs per low-power field (LPF) → Poor-quality specimen (likely saliva).

• <25 WBCs per LPF → Suggests lack of inflammation (further supports rejection).

Why Not the Other Options?

• a) Delivered within 1 hour → Ideal (prevents overgrowth of commensals).

• b) ≥25 WBCs/LPF → Good quality (indicates inflammation/infection).

• d) Early morning collection → Preferred (higher yield of pathogens).

Viral swab specimens (e.g., nasopharyngeal swabs for influenza, SARS-CoV-2) require specific transport conditions to preserve viral integrity:

1. Viral transport medium (VTM):

   • Contains stabilizers (e.g., protein, buffers) to prevent RNA/DNA degradation.

2. Refrigeration (2–8°C):

   • Slows viral degradation if testing is delayed (ideally ≤72 hours).

   • Avoid freezing (unless long-term storage is needed at –70°C or below).

Why Not the Other Options?

• a) Frozen at –20°C → Damages viral structure (ice crystals disrupt envelopes).

• c) Room temperature → Accelerates degradation (acceptable only if tested ≤24 hours).

• d) Dried on a slide → Destroys viruses (used for bacterial stains, not viral tests).

CSF is a critical, time-sensitive specimen for diagnosing meningitis and other CNS infections. The recommended handling includes:

1. Immediate transport (ideally ≤15 minutes) to the lab.

   • Delays cause false negatives due to fastidious pathogen death (e.g., Neisseria meningitidis, Streptococcus pneumoniae).

2. Room temperature storage (never refrigerate or freeze).

   • Cold temperatures kill fragile bacteria (e.g., N. meningitidis).

   • Ice can lyse cells, obscuring cytology.

Why Not the Other Options?

• a) Within 24 hours at RT → Too slow—pathogens may die.

• c) Within 2 hours, refrigerated → Refrigeration kills CSF pathogens.

• d) Within 6 hours on ice → Ice harms bacteria and cells.

Cerebrospinal fluid (CSF) must never be refrigerated for bacterial culture because:

1. Fastidious pathogens die:

   • Bacteria like Neisseria meningitidis and Streptococcus pneumoniae are extremely sensitive to cold, leading to false-negative results.

2. Cell lysis risk:

   • Refrigeration can disrupt WBCs, affecting cell counts and Gram stain accuracy.

Why Not the Other Options?

• a) Stool for Salmonella → Refrigeration is acceptable (and often recommended to prevent overgrowth of normal flora).

• b) Throat swab for Group A Streptococcus → Refrigeration is acceptable (though transport media is preferred).

• d) Urine for routine culture → Refrigeration (4°C) is standard if processing is delayed (>2 hours).

Bordetella pertussis (the causative agent of whooping cough) is fastidious and requires specific swab materials for optimal recovery:

1. Rayon or Dacron swabs:

   • Non-inhibitory to bacterial growth (unlike cotton or calcium alginate, which may contain fatty acids or residues that kill B. pertussis).

   • Compatible with PCR and culture (no interference with molecular tests).

2. Nasopharyngeal (NP) swab placement:

   • Must reach the ciliated epithelium (posterior nasopharynx) where B. pertussis colonizes.

Why Not the Other Options?

• a) Cotton swabs → May contain fatty acids that inhibit B. pertussis.

• c) Calcium alginate → Can be toxic to Bordetella and interfere with PCR.

• d) Wooden stick swabs → Not recommended (wood may contain growth inhibitors).

Minimizing contamination in blood cultures requires strict aseptic technique:

1. Two-step skin antisepsis:

   • 70% alcohol (removes surface debris) → Povidone-iodine or chlorhexidine (kills deeper flora).

   • 30-second scrub for each agent, allowing complete drying.

2. Avoiding shortcuts:

   • Never collect through IV lines (high contamination risk).

   • Multiple sets (2–4) help differentiate true bacteremia from contaminants.

Why Not the Other Options?

• a) Pre-existing IV line → Contaminated with biofilm/skin flora.

• c) Warming the hand → Helps venous access but doesn’t reduce contamination.

• d) One culture set → Inadequate (misses intermittent bacteremia; can’t rule out contamination).

• Columbia CNA (Colistin-Nalidixic Acid) agar is selective for gram-positive bacteria because:

  • CNA inhibits gram-negative bacteria (colistin disrupts membranes; nalidixic acid blocks DNA gyrase).

  • Sheep blood enriches growth of gram-positives (e.g., Staphylococcus, Streptococcus, Enterococcus).

Why Not Other Options?

• b) Trypticase soy agar with blood – Non-selective (grows gram-positive and gram-negative).

• c) Eosin methylene blue (EMB) – Selects for gram-negatives (e.g., E. coli).

• d) Modified Thayer-Martin – Selects for Neisseria gonorrhoeae (fastidious gram-negative).

False-negative blood cultures most commonly occur due to:

1. Prior antibiotic administration:

   • Antibiotics suppress bacterial growth, reducing culture sensitivity by 50–80%.

   • Even sub-therapeutic levels can inhibit fastidious organisms (e.g., Streptococcus pneumoniae).

2. Other contributing factors (but less impactful):

   • Inadequate blood volume (low pathogen load).

   • Delayed incubation (bacterial death).

Why Not the Other Options?

• b) Overfilling bottles → May cause false positives (anticoagulant dilution) but not false negatives.

• c) Proper skin disinfection → Reduces contamination, improving accuracy.

• d) Multiple sets → Increases sensitivity (detects intermittent bacteremia).

• Wooden shafts and cotton tips can contain inhibitory substances that interfere with viral growth in culture.

• Viral transport medium (VTM) is essential to maintain viral viability (dry swabs degrade viruses).

Why not other options?

• a) Incorrect – Sterile containers are required to prevent contamination.

• c) Incorrect – Antiviral therapy reduces viral load, decreasing culture sensitivity.

• d) Incorrect – Dry swabs lead to rapid viral degradation; VTM is mandatory.

Group A Streptococcus (GAS, S. pyogenes) preferentially colonizes and infects the tonsils and posterior pharynx due to:

1. Tonsillar crypts:

   • Provide a protected niche for bacterial adherence and growth.

2. High diagnostic yield:

   • Swabbing the tonsils and posterior pharynx maximizes detection of GAS (compared to oral mucosa or saliva).

Why Not the Other Options?

• a) Antibodies → Irrelevant; swabs detect bacteria, not immune responses.

• c) Free of normal flora → False; the oropharynx has abundant commensals (e.g., Viridans streptococci).

• d) Avoids saliva → Partial truth, but the primary reason is GAS tropism for tonsillar tissue.

A sterile, wide-mouth container is specifically designed for sputum collection because:

1. Ease of expectoration:

   • The wide opening allows patients to cough sputum directly into the container without spillage.

2. Minimizes contamination:

   • Sterile to avoid introducing external bacteria.

   • Secure lid prevents leakage during transport.

Why Not the Other Options?

• b) Blood for culture → Requires specialized blood culture bottles (not standard containers).

• c) Tissue biopsy → Needs anaerobic transport vials/syringes (not wide-mouth containers).

• d) Nasopharyngeal swab → Collected in viral transport media (VTM) tubes.

• Anaerobic cultures require specimens from normally sterile sites where anaerobes thrive (e.g., deep abscesses, tissue biopsies, body fluids like pleural/abdominal fluid).

• Why? Anaerobes die with oxygen exposure—specimens must be collected/transported anaerobically (e.g., in specialized tubes/swabs).

Why Not Other Options?

• a) Vaginal/eye swabs – Contaminated with normal flora (anaerobes are commensals here).

• b) Intraoral swab – Oral cavity has normal anaerobic flora (no diagnostic value).

• d) Urine/sputum – Exposed to oxygen; anaerobes are rarely pathogens here.

od cultures must be collected before antibiotics for the following critical reasons:

1. Antibiotics inhibit bacterial growth:

   • Even sub-therapeutic antibiotic levels can suppress or kill bacteria, leading to false-negative cultures.

   • This delays diagnosis and proper treatment.

2. Maximizes detection of pathogens:

   • Blood cultures rely on microbial replication for detection (e.g., by automated systems like BACTEC). Antibiotics interfere with this process.

Why Not the Other Options?

• a) Avoid contamination by skin flora → Achieved by proper skin antisepsis (e.g., alcohol/iodine), not timing of antibiotics.

• b) Prevent dilution → Unrelated; dilution depends on blood-to-broth ratio, not antibiotics.

• d) Reduce hemolysis → Hemolysis is caused by poor collection technique (e.g., vigorous mixing), not antibiotics.

Blood cultures are used to detect bloodstream infections (bacteremia/fungemia). The specimen must be collected under optimal conditions to maximize pathogen recovery.

Why Option (a) is Unacceptable:

• Antibiotics inhibit bacterial growth: If blood is drawn after antibiotics have been administered, the drugs may suppress microbial growth, leading to false-negative results.

• Best Practice: Blood cultures should ideally be collected before starting antibiotics. If antibiotics have already been given, the lab should be informed, and additional methods (e.g., antibiotic-neutralizing resins in culture bottles) may be used.

Why Other Options Are Acceptable:

• b) Blood collected from venipuncture site – This is the standard method for blood culture collection (proper skin antisepsis is required to avoid contamination).

• c) Blood collected from a sterile syringe directly into culture bottles – This is acceptable as long as proper aseptic technique is followed (no exposure to non-sterile containers).

• d) Blood collected in appropriate volume – The volume of blood is critical for sensitivity (adults typically need 8-10 mL per bottle, children 1-5 mL).

To minimize false-negative urine cultures, the most critical step is preserving specimen integrity:

1. Boric acid preservative:

   • Maintains bacterial viability for up to 24 hours at room temperature.

   • Prevents overgrowth of contaminants (e.g., Lactobacillus), which can obscure pathogens.

2. Impact on false negatives:

   • Without preservatives, fastidious bacteria (e.g., Neisseria gonorrhoeae, Streptococcus pneumoniae) may die during transport.

Why Not the Other Options?

• b) Last portion of urine → Irrelevant (midstream is standard to avoid contamination).

• c) Prolonged RT transport → Increases false negatives (bacteria die without preservatives).

• d) Non-sterile gloves → Risks contamination (but doesn’t cause false negatives).