OBJECTIVES

After completing this chapter, the reader should be able to

  • Discuss the common tests used by the microbiology laboratory for the identification of bacteria

  • List the types of clinical specimens that may be submitted for Gram stain and culture

  • Describe the process of staining and culturing a clinical specimen for bacteria, including the time required to obtain a result from either method; discuss the clinical utility of the information obtained from a Gram stain and a culture result

  • Identify bacteria according to Gram stain result (gram-positive versus gram-negative), morphology (cocci versus bacilli), and growth characteristics (aerobic versus anaerobic)

  • Define normal flora; identify anatomic sites of the human body where normal flora are commonly present and those that are usually sterile; list bacteria that are considered normal flora in each of these sites

  • Describe the most common causative pathogens based on infection type or anatomic site of infection

  • Describe the common methods used for antimicrobial susceptibility testing including technique, type of result, clinical implications, and limitations of each method; demonstrate the ability to appropriately use susceptibility information when choosing an antimicrobial regimen for a patient

  • Define minimum inhibitory concentration, MIC50, MIC90, minimum inhibitory concentration susceptibility breakpoint, and minimum bactericidal concentration

  • Describe the information that is used to construct a cumulative antibiogram; discuss the clinical utility of the cumulative antibiogram when choosing empiric antibiotic therapy for the treatment of a patient’s infection

  • List the laboratory tests that may be performed for the diagnosis of infections due to miscellaneous or uncommon organisms such as Borrelia burgdorferi, Treponema pallidum, and Legionella pneumophila

  • Describe the clinical utility of laboratory tests routinely performed for the diagnosis of infection in the following situations: (1) cerebrospinal fluid when meningitis is suspected, (2) respiratory secretions when lower respiratory tract infections are suspected, (3) urine, prostatic secretions, or genital secretions when a genitourinary tract infection is suspected, and (4) otherwise sterile fluid when infection is suspected (eg, synovial fluid, peritoneal fluid)

The assessment, diagnosis, and treatment of a patient with an infection may appear to be an overwhelming task to some clinicians. This may be partly due to the nonspecific presentation of many infectious processes; the continuously changing taxonomy, diagnostic procedures, and antimicrobial susceptibility patterns of infecting organisms; and the continuous introduction of new antimicrobials to the existing large collection of anti-infective agents. This chapter focuses on the laboratory tests used for the diagnosis of the most common infections caused by bacteria.

Information regarding white blood cells (WBCs) and their role in infection is discussed in Chapter 16; laboratory tests used in the diagnosis of viral hepatitis, Helicobacter pylori gastrointestinal (GI) infection, and Clostridioides difficile pseudomembranous colitis are addressed in Chapter 15; and lab tests used for the diagnosis of viral, fungal, and mycobacterial infections are discussed in Chapter 19. Lastly, information regarding the clinical utility of the erythrocyte sedimentation rate (ESR) and C-reactive protein (CRP) as they relate to inflammatory diseases and infections are also addressed in Chapter 20.

BACTERIA

Bacteria are small, unicellular, prokaryotic organisms that contain a cell wall but lack a well-defined nucleus. They are a diverse group of microorganisms that exist in different shapes and morphologies with varying rates of pathogenicity. Bacteria are a common cause of infection in both the community and hospital setting, and can cause infection in patients with normal or suppressed immune systems. Bacteria must be considered potential causative pathogens in any patient presenting with signs and symptoms of infection.

Specimen Collection and Identification of Bacteria

Several factors should be considered when choosing an appropriate antimicrobial regimen for the treatment of infection, including patient characteristics (eg, immune status, age, end-organ function, comorbidities, drug allergies, and severity of illness), drug characteristics (eg, spectrum of activity, pharmacokinetics, penetration to the site of infection, and proven clinical efficacy), and infection characteristics (eg, site/type of infection [suspected or known] and potential causative organism(s)). Therefore, appropriate diagnosis is a key factor in selecting appropriate empiric and directed antibiotic therapy for the treatment of an infection. In the case of a suspected infection, appropriate culture specimens should be obtained for laboratory testing from the suspected site of infection before antibiotics are initiated, if possible, in an attempt to isolate and identify the causative pathogen. Special attention should be placed on specimen collection and timely transport to the laboratory because the accuracy of the results is limited by the quality and integrity of the submitted specimen.1-4 Table 18-1 lists common biologic specimens that may be submitted to the microbiology laboratory for bacteriologic analysis.1-4

TABLE 18-1.

Common Biologic Specimens Submitted for Culture

Abscess, lesion, wound, pustule: swab or aspirate

Blood

Bone marrow

Body fluids: amniotic, abdominal, bile, pericardial, peritoneal, pleural, or synovial by needle aspiration

Bone: biopsy of infected area

CSF: by lumbar puncture or directly from shunt (eg, ventriculoperitoneal shunt)

Cutaneous: hair or nail clippings, skin scrapings, aspiration of leading edge of skin infection; biopsy

Ear: middle ear fluid specimen by myringotomy; outer ear specimen by swab or biopsy

Eye: conjunctival swab, corneal scrapings, aqueous or vitreous fluid

Foreign bodies: intravenous catheter tip by roll plate method; prosthetic heart valve, prosthetic joint material, intrauterine device, etc.

GI tract: gastric aspirate for acid fast bacilli (AFB), gastric biopsy for H pylori, rectal swab, stool cultures, stool specimen for C difficile

Genital tract: cervical, endometrial, urethral, vaginal, or prostatic secretions; ulcer biopsy

Respiratory tract: sputum, tracheal aspirate, BAL, pharyngeal or nasopharyngeal swab, sinus aspirate

Tissue: biopsy

Urine: clean-catch midstream, straight-catheterized, suprapubic aspirate

BAL = bronchoalveolar lavage.

Source: References 14.

When a specimen from the suspected site of infection is submitted to the microbiology laboratory, several microbiologic tests are performed to aid in the identification of the infecting bacteria. The most common laboratory tests used for the identification of bacteria include direct microscopic examination using specialized stains (eg, Gram stain, fluorescent stains such as acridine orange or auramine-rhodamine stains) and bacterial culture techniques to foster growth of the microorganism. Once bacteria grow in culture, additional tests are then performed to identify the infecting organism and determine susceptibility of the bacteria to various antimicrobial agents.

Gram Stain

Gram stain is the most common staining method used for the microscopic examination of bacteria and is most appropriate for the evaluation of body fluids (eg, cerebrospinal fluid [CSF], pleural, synovial, etc.), respiratory tract secretions, and wound/abscess swabs or aspirates.4 The Gram stain classifies bacteria into one of two groups, gram-positive or gram-negative, based on their reaction to an established series of dyes and decolorizers. The difference in stain uptake between gram-positive and gram-negative bacteria is primarily due to differences in their bacterial cell wall composition and permeability.5-8 Although the Gram stain does not provide an exact identification of the infecting organism (eg, Klebsiella pneumoniae versus Serratia marcescens), it does provide rapid (within minutes) preliminary information about the potential infecting organism that can be used to guide empiric antibiotic therapy while waiting for culture results, which may take 24 to 48 hours or more. The Gram stain is useful for characterizing most clinically relevant bacteria but is unable to detect intracellular bacteria (eg, Chlamydia), bacteria without cell walls (eg, Mycoplasma), and organisms that are too small to be visualized with light microscopy (eg, spirochetes).6-8

The current Gram stain methodology is a slight modification of the original process developed by Hans Christian Gram in the late 19th century.5-8 The Gram stain procedure involves several staining and rinsing steps that can be performed within a few minutes. The first step involves applying, drying, and heat-fixing a thin smear of a biological specimen to a clean glass slide. Once the slide has cooled, it is then rinsed with crystal or gentian violet (a purple dye) followed by Gram’s iodine, decolorized with an ethanol or acetone rinse, and then counterstained with safranin (a pink or red dye), with a gentle tap water rinse performed between each of these steps. The slide is then blotted dry and examined under a microscope using the oil immersion lens. If bacteria are present, they are examined for stain uptake, morphology (round = coccus, rod = bacillus), and organization (eg, pairs, clusters). Gram-positive bacteria stain purple due to retention of the crystal violet-iodine complex in their cell walls, whereas gram-negative bacteria stain red because they do not retain crystal violet and are counterstained by safranin.5,6,9 The results of the Gram stain (eg, gram-positive cocci in pairs or gram-negative rods) may provide information about the possible infecting organism before the culture results become available and, in some situations, may also be used to guide empiric antibiotic therapy. Table 18-2 lists the most likely bacteria based on Gram stain results.10,11 Once culture and susceptibility results are available, the initial empiric antibiotic regimen can be de-escalated, if necessary, to target the infecting bacteria (directed therapy).

TABLE 18-2.
Preliminary Identification of Medically Important Bacteria Using Gram Stain Results

GRAM STAIN RESULT

LIKELY BACTERIAL PATHOGEN

Gram-positive (stain purple)

 Gram-positive cocci in clusters

Staphylococcus spp.

Coagulase-positive: S aureus

Coagulase-negative: S epidermidis, S hominis, S saprophyticus, S haemolyticus, S lugdunensis, etc.

 Gram-positive cocci in pairs

Streptococcus pneumoniae

 Gram-positive cocci in chains

Viridans (α-hemolytic) streptococci (S milleri, S mutans, S salivarius, S mitis, etc.)

Group (β-hemolytic) streptococci (S pyogenes [group A], S agalactiae [group B], groups C, F, and G streptococci)

Finegoldia magna, Peptostreptococcus spp., Peptoniphilus spp., Parvimonas micra

 Gram-positive cocci in pairs and chains

Enterococcus spp. (E faecalis, E faecium, E durans, E gallinarum, E avium, E casseliflavus, E raffinosus)

Gram-positive bacilli

 Nonspore-forming

Corynebacterium spp. (C diphtheriae, C jeikeium, C striatum, etc.)

Lactobacillus spp.

Listeria monocytogenes

Cutibacterium spp.

 Spore-forming

Bacillus spp. (B anthracis, B cereus, etc.)

Clostridium spp. (C perfringens, C tetani)

Clostridioides spp. (C difficile)

Streptomyces spp.

 Branching, filamentous

Actinomyces spp. (A israelii)

Erysipelothrix rhusiopathiae

Nocardia spp. (N asteroides)

Gram-negative (stain red)

 Gram-negative cocci

Neisseria spp. (N gonorrhoeae, N meningitidis, etc.)

Veillonella spp. (V parvula)

 Gram-negative coccobacilli

Haemophilus spp. (H influenzae, H parainfluenzae, H ducreyi, etc.)

Moraxella catarrhalis

Gram-negative bacilli

 Lactose-fermenting

Aeromonas hydrophila

Citrobacter spp. (C freundii, C koseri)

Enterobacter spp. (E cloacae, E asburiae, E gergoviae, E taylorae)

Escherichia coli

Klebsiella spp. (K pneumoniae, K oxytoca, K aerogenes)

Pasteurella multocida

Vibrio cholerae

 Nonlactose-fermenting

Acinetobacter spp.

Alcaligenes spp.

Burkholderia cepacia

Morganella morganii

Proteus spp. (P mirabilis, P vulgaris)

Pseudomonas spp. (P aeruginosa, P putida, P fluorescens)

Salmonella spp. (S typhi, S paratyphi, S enteritidis, S typhimurium)

Serratia marcescens

Shigella spp. (S dysenteriae, S sonnei)

Stenotrophomonas maltophilia

 Other gram-negative bacilli

Bacteroides spp. (B fragilis, B thetaiotaomicron, B ovatus, B distasonis)

Brucella spp.

Bordetella spp.

Campylobacter jejuni

Francisella tularensis

Helicobacter pylori

Legionella spp.

Gram-variable (stain both gram-positive and gram-negative in the same smear)

 Gram-variable bacilli

Gardnerella vaginalis

Source: References 10,11.

In addition to providing a clue about the potential infecting organism, the Gram stain helps to determine the presence of bacteria in a biological specimen obtained from normally sterile body fluids (eg, CSF, pleural fluid, synovial fluid, and urine directly from the bladder) and from specimens in which infection is suspected (eg, abscess aspirate, wound swabs, sputum, and tissue); the number or relative quantity of infecting bacteria; the presence of WBCs; and the quality of the submitted specimen (eg, large numbers of epithelial cells in a sputum or urine sample may signify contamination).1,4,6,7,9

Culture and Identification

The results from the Gram stain provide preliminary information regarding the potential infecting bacteria. For the bacteria to be definitively identified, the clinical specimen is also processed to facilitate bacterial growth in culture and then observed for growth characteristics (eg, type of media, aerobic versus anaerobic, shape and color of colonies) and reactions to biochemical testing. Under normal circumstances, the results of bacterial culture are typically available within 24 to 48 hours of specimen setup and processing.

For bacteria to be grown successfully in culture, the specific nutritional and environmental growth requirements of the bacteria must be taken into consideration.3,4,8,12,13 Several clinical microbiology textbooks and reference manuals are available that can assist the microbiology laboratory with the selection of appropriate culture media and environmental conditions to facilitate the optimal growth of bacteria based on specimen type and suspected bacteria.3,4,8,12

Several types of primary culture media are available that enhance or optimize bacterial growth including nutritive media (blood or chocolate agar), differential media, selective media, and supplemental broth.1,3,8,12,13 The most commonly used bacterial growth media are listed in Table 18-3.1,3,8,12,13 Blood and chocolate agar plates are nutritive or enrichment media because they support the growth of many different types of aerobic and anaerobic bacteria. Blood agar is also considered to be differential media because it can distinguish between organisms based on certain growth characteristics, such as the differentiation between streptococci based on hemolysis patterns. MacConkey, eosin methylene blue, colistin nalidixic acid, and phenylethyl alcohol agar plates are considered selective media because they preferentially support the growth of specific organisms (eg, gram-negative or gram-positive bacteria) through the use of antimicrobials, dyes, or alcohol incorporated into their media. Trypticase soy broth and thioglycollate broth are considered supplemental media because they are used for subculturing bacteria detected on agar plates or as back-up cultures to agar plates for the detection of small quantities of bacteria in biological specimens.

TABLE 18-3.
Commonly Used Bacterial Growth Media

GROWTH MEDIUM

COMPOSITION

USES

Agars

 Blood agar, SBA

5% sheep blood

The most commonly used all-purpose medium with ability to grow most bacteria, fungi, and some mycobacteria; also used for determination of hemolytic activity of streptococci

 Chocolate agar, enriched

2% hemoglobin or Iso-VitaleX in peptone base

All-purpose medium that supports growth of most bacteria; especially useful for growth of fastidious bacteria, such as Haemophilus spp., Brucella spp., and pathogenic Neisseria spp.

 EMB or Mac agar

Peptone base with sugars and dyes that yield differentiating biochemical characteristics

Included in primary setup of nonsterile specimens; selective isolation of gram-negative bacteria; differentiates between lactose-fermenting and nonlactose-fermenting enteric bacteria

 PEA or CNA agar

Nutrient agar bases with supplemental agents to inhibit growth of aerobic gram-negative bacteria

Included in primary setup of nonsterile specimens; selective isolation of gram-positive cocci and bacilli as well as anaerobic gram-positive cocci or gram-negative bacilli

Broths

 TSB

All-purpose enrichment broth

Used for subculturing bacteria from primary agar plates; supports the growth of many fastidious and nonfastidious bacteria

 Thioglycollate broth

Pancreatic digest of casein, soy broth, and glucose

Supports the growth of aerobic, anaerobic, microaerophilic, and fastidious bacteria

CNA = colistin-nalidixic acid; EMB = eosin methylene blue; Mac = MacConkey; PEA = phenylethyl alcohol; SBA = sheep blood agar; TSB = trypticase soy broth.

Source: References 1,3,8,12,13.

Once a clinical specimen is processed on growth media, the plates must be incubated in the appropriate environment to support bacterial growth. The environmental factors that should be controlled during incubation include oxygen or carbon dioxide content, temperature, pH, and moisture content of the medium and atmosphere.1,12,13 The oxygen requirements for growth differ among organisms. Strict aerobic bacteria, such as Pseudomonas aeruginosa, grow best in ambient air containing 21% oxygen and a small amount of carbon dioxide.1 Strict anaerobes, such as Bacteroides spp., are unable to grow in an oxygen-containing environment and require a controlled environment containing 5% to 10% carbon dioxide for optimal growth. Facultative anaerobes, such as Escherichia coli and some streptococci, can grow in the presence or absence of oxygen. Overall, most clinically relevant bacteria grow best at 35°C to 37°C (the temperature of the human body), with a pH of 6.5 to 7.5, and in an atmosphere rich in moisture, which is why agar plates are sealed (to trap moisture) and humidified incubators are used.12 Bacteria grown successfully in culture appear as colonies on the agar plates.

Bacterial identification is based on the results of genotypic and phenotypic testing.12 Genotypic bacterial identification tests use molecular techniques for the detection of a particular gene or RNA product that is characteristic of specific bacteria. Phenotypic bacterial identification tests involve the observation of the physical and metabolic properties of a bacteria, including the evaluation of colony characteristics (size, pigmentation, shape, and surface appearance); the assessment of culture media and environmental conditions that supported bacterial growth; the changes that occurred to the culture media as a result of bacterial growth; the aroma of the bacteria; the Gram stain result of individual colonies; and the results from biochemical testing.3,12 Biochemical tests for bacterial identification are either enzyme based, in which the presence of a specific enzyme is measured (eg, catalase, oxidase, indole, or urease tests), or based on the presence and measurement of metabolic pathways or byproducts (eg, oxidative and fermentation tests or amino acid degradation).3,12 Examples of biochemical tests include the presence of catalase in the organism or the ability of a bacteria to ferment glucose. Most biochemical tests are performed using manual or automated commercial (preferred) identification systems.12,14 Some of the commercial identification systems consist of multicompartment biochemical tests in a single microtiter tray so that several biochemical tests can be performed simultaneously.12,14 Information derived from the macroscopic examination of the bacteria and the results of biochemical tests are then combined to determine the specific identity of the bacteria. Using traditional methodology, bacterial identification is usually achieved within 24 to 48 hours of detection of bacterial growth.12,13

Numerous rapid diagnostic tests are available for identification of bacteria (with some tests also detecting several pertinent bacterial resistance genes) directly from clinical specimens such as blood, stool, respiratory secretions, or body fluids (ie, saliva, urine, CSF) using a variety of methodologies that have all been designed to substantially decrease the time to organism identification (with results in 15 minutes to 12 hours) when compared with traditional bacterial culture and identification methods.3,11,14,15 Rapid diagnostic tests are highly sensitive and specific and have become important tools for antimicrobial stewardship programs because they allow for quicker antimicrobial de-escalation/discontinuation and implementation of infection control procedures, such as isolation, when necessary.3,11,14,15 Rapid diagnostic tests are usually more costly than traditional bacterial identification methods; however, when they are used in conjunction with antimicrobial stewardship programs, the cost-effectiveness of many of the tests have been justified through faster optimization of antimicrobial therapy, improved patient outcomes, and overall lower hospital costs.15,16

The currently available rapid bacterial identification tests are immunologically based, nucleic acid (NA) based (nonamplified and amplified), or proteomic based.3,12,14,16 Immunologic methods use immunofluorescent or enzyme-linked immunosorbent assay antigen or antibody detection. Immunologically based rapid bacterial identification tests are available for the detection of group A streptococcus (pharyngeal), Streptococcus pneumoniae (urine, respiratory), Neisseria gonorrhoeae (urethral, cervical), Neisseria meningitidis (CSF), L. pneumophila (urine), influenza/respiratory syncytial virus (RSV) (nasopharyngeal), and other viruses (see respective sections within this chapter and Chapter 19 for additional information).

Some commonly used nonamplified bacterial identification methods using NA probes include peptide NA fluorescence in situ hybridization (PNA-FISH; PNA FISH and QuickFISH [Opgen, Gaithersburg, MD]), and Verigene (Luminex, Austin, TX).3,14,16 PNA-FISH and QuickFISH detect species-specific RNA using fluorescent probes and can differentiate between commonly encountered bacteria from positive blood cultures after Gram stain. PNA-FISH and QuickFISH kits are available to differentiate between Staphylococcus aureus and coagulase-negative staphylococci from blood culture bottles with gram-positive cocci in clusters; Enterococcus faecalis versus Enterococcus faecium versus other enterococci from blood cultures positive for gram-positive cocci in pairs and chains; E coli, K pneumoniae versus P aeruginosa from blood cultures positive for gram-negative rods; and Candida albicans/Candida parapsilosis versus Candida krusei/Candida glabrata from blood cultures positive for yeast. PNA-FISH results are available within 90 minutes whereas QuickFISH results are available within 20 minutes of blood culture positivity.

Verigene (Luminex, Austin TX) is a novel microarray, multiplex test using NA extraction and array hybridization in a conserved genetic region of the bacteria or virus to differentiate among different species. Several Verigene tests are currently available for the rapid identification of bacteria (and viruses) causing infection in the bloodstream, respiratory tract, and GI tract. The Verigene Gram-Positive Bloodstream Infection Test (Luminex) can identify 13 different gram-positive bacterial species and the presence of three notable gram-positive resistance gene markers, namely mecA (methicillin-resistance), vanA/vanB (vancomycin resistance) within 2.5 hours of blood culture positivity.3,16 The Verigene Gram-Negative Bloodstream Infection Test (Luminex) can identify nine different gram-negative bacteria and the presence of six resistance gene markers (CTX-M β-lactamase and several carbapenemases) within 2 hours of blood culture positivity. Verigene tests are also available for rapid pathogen identification (within 2 hours) of respiratory tract infections (Verigene Respiratory Pathogens Flex Test with three bacterial and 13 viral targets); GI infections (Verigene Enteric Pathogens Test with five bacterial, two viral, and two toxin targets); and C difficile (with the ability to detect the presence of toxin A, toxin B, and polymerase chain reaction [PCR] ribotype 027 hypervirulent strain).

Multiple amplified NA-based tests, including PCR and other methods, are available for the rapid identification of bacteria (and occasionally the presence of resistance gene markers) and viruses.16 Currently, the most comprehensive test is the FilmArray multiplex PCR system (BioFire Diagnostics, Salt Lake City, UT), which integrates sample preparation, amplification, detection, and analysis into one process with results available within 1 hour.3,16 FilmArray is available in several U.S. Food and Drug Administration (FDA)–approved panels for the quick identification of the causative pathogens of meningitis/encephalitis, bloodstream infections, respiratory tract infections, and GI tract infections (https://www.biofiredx.com/products/filmarray/).

Lastly, matrix-assisted laser desorption-ionization time-of-flight (MALDI-TOF) mass spectrometry is a proteomic-based rapid pathogen identification test that has become widely used by many clinical microbiology labs because it is an inexpensive test that provides the quick identification of a large number of clinically relevant pathogens.3,14 The bacteria are placed on a target plate with a matrix solution and then pulsed with a laser. The mass of the ionized particles produced by the laser differx based on the organism, and the pathogen and relative quantity within the sample can be identified within 10 to 30 minutes using a library of standard reference species.14,16

Colonization, Contamination, or Infection

The growth of an organism from a submitted biologic specimen does not always indicate the presence of infection; it may represent the presence of colonization or contamination.9,11 Table 18-4 lists the anatomic sites, fluids, and tissues of the human body that are sterile, including the bloodstream, the CSF, internal organs and tissues, bone, synovial fluid, peritoneal fluid, pleural fluid, pericardial fluid, and urine taken directly from the bladder or kidney. Alternatively, other body sites, particularly those with a direct connection to the outside environment, are naturally colonized with microorganisms called normal flora. Normal bacterial flora can be found on the skin, in the oral cavity, and in the respiratory, GI, and genitourinary tracts and may be recovered from clinical specimens obtained from these areas. Examples of bacteria that typically colonize these body sites are listed in Table 18-5.11,17 Typically, normal flora are harmless bacteria that rarely cause infection. They are often located in the same areas of the body as pathogenic bacteria and are thought to provide protection by inhibiting the growth of pathogenic organisms through competition for nutrients and by stimulating the production of cross-protective antibodies.11,17 However, normal flora may potentially become pathogenic and cause infection in patients with suppressed immune systems or after translocation to normally sterile body sites during trauma, intravascular line insertion, or surgical procedures, especially when the skin is not adequately cleansed in the latter two situations. Alternatively, pathogenic bacteria may occasionally colonize body sites, where they are present but do not invade host tissue or elicit signs and symptoms of infection (Table 18-6).

TABLE 18-4.

Normally Sterile Body Sites

Bloodstream

CSF

Pericardial fluid

Pleural fluid

Peritoneal fluid

Synovial fluid

Bone

Urine (directly from the bladder or kidney)

TABLE 18-5.

Body Sites with Normal Colonizing Bacterial Flora

SKIN

ORAL CAVITY

Corynebacterium spp.

Cutibacterium spp.

Staphylococcus spp. (especially coagulase-negative staphylococci)

Streptococcus spp.

Actinomyces spp.

Prevotella spp.

Viridans streptococci

Anaerobic streptococci

Veillonella spp.

RESPIRATORY TRACT

GI TRACT (COLON)

Viridans streptococci

Anaerobic streptococci

Haemophilus spp.

Neisseria spp.

Moraxella spp.

Bacteroides spp.

Clostridium spp.

Escherichia coli

Klebsiella pneumoniae

Enterococcus spp.

Anaerobic streptococci

GENITOURINARY TRACT

Lactobacillus spp.

Streptococcus spp.

Staphylococcus spp.

Corynebacterium spp.

Enterobacterales

Mycoplasma hominis

Bacteroides spp.

Prevotella spp.

Source: References 11,17.
TABLE 18-6.

Clinical, Laboratory, and Radiographic Signs and Symptoms of Infection

CLINICAL

Localized

 Pain and inflammation at site of infection: erythema, swelling, warmth

 Purulent discharge (wound, vaginal, urethral discharge)

 Sputum production and cough (pneumonia)

 Diarrhea

 Dysuria, frequency, urgency, suprapubic tenderness, costovertebral angle tenderness (UTI)

 Headache, nuchal rigidity, photophobia, Brudzinski’s sign, Kernig’s sign (meningitis)

Systemic

 Fever

 Chills, rigors

 Malaise

 Tachycardia

 Tachypnea

 Hypotension

 Mental status changes

LABORATORY

 Increased WBC count: peripherally and/or at the site of infection

Decreased WBC count: occasionally, patients present with leukopenia

 Increased neutrophil percentage, including an increase in immature neutrophils (bands or stabs) in the WBC differential called a “shift to the left”

 Hypoxemia (pneumonia)

 Elevated lactate

 Positive Gram stain and/or culture from site of infection

 Elevated ESR and CRP

 Elevated procalcitonin levels

 Positive antigen or antibody test, positive PCR test, elevated antibody titers

RADIOGRAPHIC

 Chest radiograph with consolidation, infiltrate, effusion, or cavitary nodules (lung infection)

 Bone radiograph or MRI: periosteal elevation or bone destruction (osteomyelitis)

 CT/MRI: rim-enhancing lesions (abscess)

CT = computed tomography; MRI = magnetic resonance imaging.

Contamination occurs when an organism is accidentally introduced into a biologic specimen during specimen collection, transport, or processing.1,3,4,9,11 Bacteria that cause contamination typically originate from the skin of the patient (especially if the skin is not adequately cleaned before specimen acquisition), the clinician, or the laboratory technician but may also come from the environment. The most common contaminant is coagulase-negative staphylococci (especially Staphylococcus epidermidis), which is an organism that normally colonizes the skin.17 Other common contaminants of blood culture specimens include Micrococcus spp., Cutibacterium acnes, and most Bacillus and Corynebacterium spp.17

Infection occurs when an organism is present and invades or damages host tissues, eliciting a host response and producing signs and symptoms consistent with an infectious process. When determining the presence of infection in an individual patient, several factors should be considered, such as the clinical condition of the patient (eg, fever and purulent discharge), the presence of laboratory signs of infection (eg, high WBC count), and the results of radiographic and microbiologic tests.18 Table 18-6 describes some of the local and systemic clinical signs and symptoms, laboratory findings, and radiographic findings that may be present in a patient with infection. The exact clinical, laboratory, and radiographic signs of infection differ based on the site of infection, the age of the patient, and the severity of the infection. For example, a patient with pneumonia usually exhibits a fever, productive cough, shortness of breath, tachypnea, leukocytosis, and an infiltrate on chest radiograph, whereas a patient with an uncomplicated urinary tract infection (UTI) typically experiences urinary frequency, urgency, dysuria, and pyuria. It is important to note that the typical signs and symptoms of infection may not be present in elderly persons or in patients who are immunocompromised (eg, patients with neutropenia and patients with acquired immune deficiency syndrome).

False-positive culture results can lead to the use of additional laboratory tests, radiographic tests, unnecessary antibiotics, increased length of hospitalization, and patient costs; therefore, every positive culture should warrant an evaluation for clinical significance. The diagnosis of infection should be suspected in any patient with a positive culture accompanied by clinical, laboratory, and radiographic findings suggestive of infection. Luckily, certain bacteria have a propensity to cause infection in particular body sites and fluids, as demonstrated in Table 18-7.4,18,19 This information can be used to determine if the bacteria isolated from a culture is a commonly encountered pathogen at the particular site of infection.8 For instance, the growth of S pneumoniae from the sputum of a patient with signs and symptoms consistent with community-acquired pneumonia (CAP) is a significant finding because S pneumoniae is the most common cause of CAP. However, the growth of S epidermidis from a blood or wound culture from an asymptomatic patient should be evaluated for clinical significance because it may represent contamination of the submitted specimen.11,18 The information in Table 18-7 regarding the most common causative organisms by infection site can also be used to select empiric antibiotic therapy before culture results are available by guiding the selection of an antibiotic regimen with activity against the most common causative bacteria at the suspected site of infection, as illustrated in Minicase 1.19

TABLE 18-7.

Common Pathogens by Site of Infection

MOUTH

SKIN AND SOFT TISSUE

BONE AND JOINT

Anaerobic streptococci

Peptococcus spp.

Peptostreptococcus spp.

Actinomyces israelii

Staphylococcus aureus

Streptococcus pyogenes

Staphylococcus epidermidis

Pasteurella multocida

Clostridium spp.

Staphylococcus aureus

Streptococcus pyogenes

Streptococcus spp.

Staphylococcus epidermidis

Neisseria gonorrhoeae

Gram-negative bacilli

INTRA-ABDOMINAL

URINARY TRACT

UPPER RESPIRATORY TRACT

Escherichia coli

Proteus mirabilis

Klebsiella spp.

Enterococcus spp.

Bacteroides spp.

Clostridium spp.

Escherichia coli

Proteus mirabilis

Klebsiella spp.

Enterococcus spp.

Staphylococcus saprophyticus

Streptococcus pneumoniae

Haemophilus influenzae

Moraxella catarrhalis

Streptococcus pyogenes

LOWER RESPIRATORY TRACT, COMMUNITY-ACQUIRED

LOWER RESPIRATORY TRACT, HOSPITAL-ACQUIRED

MENINGITIS

Streptococcus pneumoniae

Haemophilus influenzae

Moraxella catarrhalis

Klebsiella pneumoniae

Legionella pneumophila

Mycoplasma pneumoniae

Chlamydia pneumoniae

Early onset (within 4 days of hospitalization)

Klebsiella pneumoniae

Escherichia coli

Enterobacter spp.

Proteus spp.

Serratia marcescens

Haemophilus influenzae

Streptococcus pneumoniae

 MSSA

Late onset (> 4 days after hospitalization)

 Pathogens above plus MDR organisms:

Acinetobacter spp.

Pseudomonas aeruginosa

 MRSA

Legionella pneumophila

Streptococcus pneumoniae

Neisseria meningitidis

Haemophilus influenzae

Group B Streptococcus

Escherichia coli

Listeria monocytogenes

MDR = multidrug resistant; MRSA = methicillin-resistant Staphylococcus aureus; MSSA = methicillin-susceptible Staphylococcus aureus.

Source: References 4,18,19.

Occasionally, culture results may be negative in patients with infection, particularly in the setting of previous antibiotic use, improper specimen collection, or the submission of inadequate specimens. In this setting, the clinical condition of the patient may establish the presence of infection despite negative cultures, and the suspected site of infection should help guide antibiotic therapy based on most likely causative organisms that typically cause infection at that site.9

Antimicrobial Susceptibility Testing

Once an organism has been cultured from a biologic specimen, further testing is performed in the microbiology laboratory to determine antibiotic susceptibility of the infecting organism to help direct and streamline antimicrobial therapy. Because of the continued emergence of resistance in some bacteria, susceptibility testing is imperative for determining the antimicrobial agents that should be used for the treatment of the patient’s infection. Several methods are available for determining antibiotic susceptibility of a particular organism by either directly measuring the activity of an antibiotic against the organism or detecting the presence of a specific resistance gene/mechanism in the organism, as described in Table 18-8.9,11,20-25 Microbiology laboratories often use several different methods for susceptibility testing to accurately determine the activity of antibiotics against many different types of bacteria (eg, aerobic, anaerobic, and fastidious). The Clinical and Laboratory Standards Institute (CLSI) continuously updates and publishes standards and guidelines for the susceptibility testing of aerobic and anaerobic bacteria to assist microbiology laboratories in determining the specific antibiotics and test methods that should be used for particular organisms or specific clinical situations/infections.26-28

TABLE 18-8.

Antimicrobial Susceptibility Testing Methods

Methods that directly measure antibiotic activity

Dilution susceptibility tests: broth macrodilution (tube dilution), broth microdilution, agar dilution

Disk diffusion: Kirby-Bauer

Antibiotic concentration gradient methods: Etest

Other specialized tests:

  Measure bactericidal activity: MBC testing, time-kill studies, SBT

  Susceptibility testing of antibiotic combinations (synergy testing): checkerboard technique, time-kill curve technique, disk diffusion, Etest

Methods that detect the presence of antibiotic resistance or resistance mechanisms

 β-lactamase detection

 Detection of HLAR

 Agar screens for detection of MRSA or VRE

 Chloramphenicol acetyltransferase detection

 Molecular methods involving NA hybridization and amplification

 Penicillin-binding protein (PBP) 2a

 Inducible clindamycin resistance

Etest = epsilometer test; HLAR = high-level aminoglycoside resistance; MBC = minimum bactericidal concentration; NA = nucleic acid; SBT = serum bactericidal tests; VRE = vancomycin-resistant enterococci.

Source: References 9,11,2025.

Methods That Directly Measure Antibiotic Activity

Several tests directly measure the activity of an antibiotic against a particular organism. Quantitative tests measure the exact concentration of an antibiotic necessary for inhibiting the growth of the bacteria whereas qualitative tests measure the comparative inhibitory activity of several antibiotics against the organism. The format of the reported test results and interpretation of susceptibility from each of these methods is different depending on the methodology used. The advantages and disadvantages of the different antimicrobial susceptibility testing methods are listed in Table 18-9.9,11,20,25,29,30

TABLE 18-9.

Advantages and Disadvantages of Antimicrobial Susceptibility Testing Methods

METHOD

ADVANTAGES

DISADVANTAGES

Agar dilution

An exact MIC is generated

Several isolates can be tested simultaneously on the same plate at a relatively low cost

Susceptibility of fastidious bacteria can be determined because agar supports their growth

Time-consuming

Antibiotic plates need to be prepared manually when needed and can be stored only for short periods of time

Plates are not commercially available; must be prepared by the laboratory

Broth macrodilution (tube dilution)

An exact MIC is generated

The MBC can also be determined, if desired

Each antibiotic is tested individually

Method is labor and resource intensive

Broth microdilution (automated)

Simultaneously tests several antibiotics

Less labor and fewer resources are used

Commercially prepared trays or cards can be used

MIC range (rather than exact MIC) is typically reported

The number of antibiotics and concentrations that are tested are predetermined and limited

Disk diffusion (Kirby-Bauer)

Simultaneously tests several antibiotics

Exact MICs cannot be determined

Cannot be used for fastidious or slow-growing bacteria

Etest

An exact MIC is generated

Easy to perform

Several antibiotics can be tested on the same plate

Relatively expensive

Not all antibiotics are available as Etest strips

Etest = epsilometer test; MIC = minimum inhibitory concentration.

Source: References 9,11,20,25,29,30.

Choosing Empiric Antibiotic Therapy for Hospital-Acquired Pneumonia

Marie A., a 68-year-old woman with no known drug allergies, is admitted to the general medical floor of University Hospital for management of a right cerebral vascular accident after stabilization in the emergency department. Prior to admission, she was living at home with her husband and was previously healthy without recent hospitalizations or antibiotic therapy within the past few years. She continues to have left-sided hemiparesis and has been deemed to be an aspiration risk by physical therapy/occupational therapy. On hospital day 3, she develops a temperature of 102.3°F, chills, tachypnea, a productive cough, and shortness of breath requiring supplemental oxygen via nasal cannula. Her physical exam reveals an increased respiration rate (RR) of 24 breaths/min and decreased breath sounds in the right middle lobe. Her laboratory results reveal a total WBC count of 18,000 cells/mm3 with 70% neutrophils, 19% bands, 7% lymphocytes, and 4% monocytes. Her chest radiograph demonstrates right middle lobe consolidation consistent with pneumonia. An expectorated sputum sample is obtained for Gram stain and culture. The Gram stain reveals >25 WBC/hpf, <10 epi/hpf, and many gram-negative rods. Her physician asks you to recommend empiric antibiotic therapy to treat her pneumonia before the final culture results are available.

QUESTION: What is the most likely causative organism of this patient’s pneumonia, and which empiric antibiotic therapy would you choose based on the Gram stain results?

DISCUSSION: This patient most likely has early-onset (within 4 days of hospitalization) HAP, in which the most common causative organisms (Table 18-7) include S pneumoniae; H influenzae; gram-negative bacteria such as K pneumoniae, E coli, Enterobacter spp., S marcescens, and Proteus spp.; S aureus (methicillin-susceptible Staphylococcus aureus); and atypical bacteria such as L pneumophila (especially in patients with diabetes mellitus, underlying lung disease, renal failure, or suppressed immune systems). Based on the Gram stain results (Table 18-2) demonstrating the presence of gram-negative rods (not unexpected because gram-negative bacteria are the most common cause of HAP overall), the most likely causative organisms include K pneumoniae, E coli, Enterobacter spp., S marcescens, or Proteus spp. Based on the most recent Infectious Diseases Society of America guidelines for the management of HAP, the patient should receive empiric therapy with piperacillin–tazobactam, cefepime, or a fluoroquinolone (levofloxacin) because she is not at high risk for mortality from HAP (not on ventilatory support or in septic shock) and does not have risk factors for a multidrug-resistant organism.19 The antibiotic regimen can be modified to more directed therapy, if possible, once the results of the culture and susceptibility are available.

Dilution Methods (Macrodilution and Microdilution)

Several dilution methods measure the activity of an antibiotic against a particular organism. Both broth dilution and agar dilution methods quantitatively measure the in vitro activity of antibiotics against a particular organism, with results reported as the minimum inhibitory concentration (MIC). Broth dilution can be performed using macrodilution or microdilution, in which the main differences between the methods include the volume of broth used, the number of antibiotics that can be simultaneously tested, and the manner in which the test results are generated and reported. The agar dilution method differs in that it is performed using solid growth media.

Broth macrodilution

Broth macrodilution, or the tube-dilution method, is one of the oldest methods of antimicrobial susceptibility testing and is often considered the gold standard. This method is performed in test tubes containing 2-fold serial dilutions of the antibiotic being tested for susceptibility (with concentrations typically representing clinically achievable serum or site concentrations of the antibiotic in mcg/mL) in a liquid growth media (1 mL of broth or greater) to which a standard inoculum (5 × 105 CFU/mL) of the infecting bacteria is added.9,20,25-27,29 The test tubes are incubated for 16 to 24 hours at 35°C and then examined macroscopically for the presence of turbidity or cloudiness, which is an indication of bacterial growth.11,20,25,27,29 The test tube containing the lowest antibiotic concentration that completely inhibits visible growth (the broth in the tube appears clear to the unaided eye) represents the MIC in mcg/mL (Figure 18-1).9,20,25,27,29

FIGURE 18-1.
FIGURE 18-1.
Broth macrodilution susceptibility testing for MIC and MBC. (Source: Reprinted with permission from Graman PS, Menegus MA. Microbiology laboratory tests. In: Betts RF, Chapman SW, Penn RL, eds. A Practical Approach to Infectious Diseases. 5th ed. Philadelphia, PA: Lippincott, Williams and Wilkins; 2003:929–956.)

The CLSI has established interpretive criteria for MIC results of each antibiotic against each bacteria as susceptible (S), susceptible-dose dependent (SDD), intermediate (I), and resistant (R). The MIC value that separates or defines these categories for an antibiotic are known as MIC breakpoints.20,21,25,26 MICs have been categorized as S, SDD, I, and R to help predict the probable response of a patient’s infection to a particular antibiotic.9,21,25,26 Bacteria that are categorized as susceptible to a given antibiotic will, most likely, be eradicated during treatment of the infection because concentrations of the antibiotic represented by the MIC are easily achievable using standard doses of the antibiotic. Susceptible-dose dependent is a category applied to specific antibiotic-organism pairs (eg, cefepime and Enterobacterales) in which several approved or routinely used dosing options are available. Bacteria that are categorized as SDD require treatment with higher and/or more frequent doses (usually higher doses than were used when establishing the S breakpoint) to achieve higher drug exposures for treatment of the infection. Intermediately susceptible bacteria display higher MICs, and successful treatment may be achieved if higher than normal doses of an antibiotic are used or if the antibiotic concentrates at the site of infection.25 In clinical practice, antibiotics that display intermediate susceptibility are rarely used for treatment of infection because clinical response is unpredictable; however, they may be considered for treatment when the organism displays resistance to all other agents tested. Lastly, organisms that are resistant to an antibiotic display extremely high MICs that exceed the normal achievable serum concentrations of the antibiotic, even if maximal doses are used, resulting in a poor clinical response. In general, it is the responsibility of the clinician to determine if a drug listed as susceptible from an individual isolate susceptibility report is useful for the treatment of a particular infection based on the pharmacokinetic parameters (site penetration) and clinical efficacy studies of the antibiotic for that infection type.

Minimum inhibitory concentration breakpoints for each antibiotic and bacteria are different because they are based on achievable serum concentrations of the antibiotic with normal dosing; the inherent susceptibility of the organism to the antibiotic; the site of infection and ability of the antibiotic to obtain adequate concentrations at that site; pharmacodynamic analysis with Monte Carlo simulations to predict efficacy; and the results of antibiotic clinical trials evaluating efficacy based on organism MIC.20,21,26,27 The safe and effective dose of each antibiotic is typically determined using pharmacokinetic, safety, and efficacy data gathered during preclinical stages of drug development. Because each antibiotic has its own unique pharmacokinetic profile and recommended dosage range, it is not surprising that each antibiotic achieves different serum concentrations after standard dosing. For example, intravenously administered piperacillin–tazobactam achieves higher serum concentrations and area under the serum concentration time curve than intravenously administered levofloxacin. Therefore, the MIC susceptibility breakpoint for piperacillin–tazobactam against Enterobacterales is higher (≤16 mcg/mL) than levofloxacin (≤0.5 mcg/mL).26

As mentioned previously, there are other factors to consider when MIC breakpoints are established. Some antibiotics are inherently more active against an organism than others, which is reflected by a lower MIC required to inhibit bacterial growth. The site of infection may predict the potential usefulness of the antibiotic depending on its ability to achieve adequate concentrations at the site of infection. An antibiotic might display potent in vitro activity against a particular organism but may be ineffective in vivo due to poor penetration to the site of infection. In fact, there are a few clinical situations in which the site of infection is directly incorporated into the MIC susceptibility interpretation of an antibiotic, such as in the case of meningitis caused by S pneumoniae in which the interpretation of ceftriaxone and penicillin susceptibility are determined using meningitis MIC breakpoints (which are lower).26 Monte Carlo simulations may also be performed to help establish MIC breakpoints. Analyses are performed using population pharmacokinetic data of the antibiotic and bacteria MIC distribution data from susceptibility studies to evaluate the percentage of time the antibiotic will achieve adequate pharmacodynamic indices for the treatment of that organism in a simulated population. Lastly, results from clinical trials evaluating antibiotic efficacy are also considered in MIC breakpoint determination where a correlation is made between clinical efficacy and the MIC value of the infecting organism. For example, what MICs were observed in patients with clinical success versus clinical failure?

Broth macrodilution is useful because an exact MIC of the infecting organism can be derived. The results of broth macrodilution are reported as the MIC of the antibiotic against the infecting organism with its corresponding interpretive category (S, I, and R). However, broth macrodilution is rarely used in microbiology laboratories because the methodology is resource and labor intensive, making it impractical for everyday use.

An additional step can be performed on the broth macrodilution test to determine the actual antibiotic concentration that kills 99.9% of the bacterial inoculum, which is also known as the minimum bactericidal concentration (MBC).20,22,24 Samples from all of the test tubes that did not exhibit visible growth are subcultured on agar plates and incubated at 35°C for 18 to 24 hours (Figure 18-1).9,22 The plate representing the lowest antibiotic concentration that does not support the growth of any bacterial colonies is defined as the MBC. Because a higher concentration of an antibiotic may be necessary to kill the organism rather than just inhibit its growth, the MIC will be equal to or lower than the MBC. The determination of the MBC is not routinely performed in clinical practice and may be considered in rare clinical circumstances, such as during the treatment of severe or life-threatening infections (eg, endocarditis, meningitis, osteomyelitis, or sepsis in immunocompromised patients) in patients who are not responding to therapy.9,22,24

Broth microdilution

Broth microdilution susceptibility testing was developed to overcome some of the limitations of the broth macrodilution method and has become the most commonly used method for susceptibility testing of bacteria in microbiology laboratories.11,20,25,27,29 Instead of using standard test tubes with 2-fold serial dilutions of antibiotics, this method uses manually prepared or commercially prepared disposable microtiter cassettes or trays containing up to 96 wells that can simultaneously test the susceptibility of up to 12 antibiotics depending on the product used.11,20,25,27,29 Several examples of microtiter trays are shown in Figure 18-2A and B.20 The wells in the broth microdilution trays contain a smaller volume of broth (0.05 to 0.1 mL) to support bacterial growth than broth macrodilution (1 mL or more). The microtiter tray is inoculated with a standardized inoculum of the infecting organism and incubated for 16 to 20 hours. The tray is then examined for bacterial growth by direct visualization using light boxes, direct visualization using reflecting mirrors, or automated, computer-assisted readers. The MIC represents the microdilution well containing the lowest antibiotic concentration that completely inhibits visible bacterial growth (eg, did not produce turbidity). Several companies commercially supply broth microdilution panels that contain broth with appropriate antibiotic concentrations according to guidelines for conventional broth dilution methods. Some examples of manual/semiautomated systems include Sensititre OptiRead or Vizion System (Thermo Fisher Scientific, Waltham, MA) and MicroScan autoSCAN-4 (Beckman Coulter, Brea, CA). Examples of fully automated systems include Vitek 2 or Vitek-Legacy (bioMérieux Diagnostics, Marcy-l’Étoile, France), MicroScan WalkAway plus System (Beckman Coulter), Sensititre ARIS 2X (Thermo Fisher Scientific), and the Phoenix Automated Microbiology System (BD, Franklin Lakes, NJ).20 Some of these systems also have been designed to identify bacteria and are able to provide more rapid susceptibility results (within 8 hours) due to shortened incubation times.

FIGURE 18-2.
FIGURE 18-2.
(A) Broth microdilution susceptibility panel containing 96 reagent wells and a disposable tray inoculator. (B) Example of the Vitek 2 antimicrobial susceptibility test card with 64 wells with multiple concentrations of up to 22 antibiotics. (Source: Reprinted with permission from Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:177–204.)

Because of the size constraints of the broth microdilution panels, only a limited number of antibiotics and concentrations can be incorporated into the trays. Typically, drugs that have inherent activity against the class of bacteria being tested (eg, gram-positive versus gram-negative) are included in the trays. For example, when determining the susceptibility of gram-negative bacteria, it is impractical to include antibiotics in the microdilution trays that do not have activity against these organisms, such as penicillin, nafcillin, or vancomycin. The same holds true for susceptibility testing of gram-positive organisms, where it would be impractical to test the susceptibility of piperacillin or ceftazidime against S aureus because these agents have limited antistaphylococcal activity. In addition, the trays are not large enough to incorporate the full range of antibiotic concentrations usually tested during broth macrodilution. Therefore, the concentrations incorporated into the wells for each antibiotic often reflect the CLSI interpretive category breakpoints of S, I, and R for the particular antibiotic-organism combination.

The results of broth microdilution tests are reported as either an exact MIC or MIC range with corresponding CLSI interpretation. In this case, an MIC range may be reported due to the limited antibiotic concentrations tested for each antibiotic. For example, if bacterial growth is not detected in the lowest concentration tested of a particular antibiotic using broth microdilution, the MIC would be reported as less than or equal to that concentration tested. The MIC could be much lower, but the exact MIC could not be determined because lower concentrations of the antibiotic were not tested because of the size constraints of the microdilution trays.

The advantages of broth microdilution include the ability to test the susceptibility of multiple antibiotics simultaneously, the ease of use when commercially prepared microdilution trays are used, rapid results with the automated methods, and decreased cost/labor.20,25,27 The disadvantages of broth microdilution include the lack of flexibility of antibiotics available in commercially prepared microdilution trays, the limitation on the number of concentrations that can be tested for each antibiotic due to size constraints of the trays, and the reporting of an MIC range (on many occasions) rather than the true MIC against an infecting organism.20,25

Agar dilution

Agar dilution is another quantitative susceptibility testing method that uses 2-fold serial dilutions of an antibiotic incorporated into agar growth medium, with each concentration placed into individual Petri dishes.20,25,29 The surface of each plate is inoculated with a standardized bacterial inoculum (1 × 104 CFU/mL) and incubated for 18 to 20 hours at 35°C. The susceptibility of several different bacteria can be evaluated simultaneously on the plates. The MIC is represented by the plate with the lowest concentration of antibiotic that does not support visible growth of the bacteria. The advantages of agar dilution include the ability to simultaneously test the susceptibility of several different bacteria, the ability to perform susceptibility testing of fastidious organisms given agar adequately supports their growth, and the generation of an exact MIC of the infecting bacteria. However, agar dilution is not commonly used in most microbiology laboratories because it is resource and labor intensive. In addition, the antibiotic plates are not commercially available and must be prepared before each susceptibility test because they can only be stored for short periods of time.20,25,29

Disk Diffusion Method (Kirby-Bauer)

The disk diffusion method is a well-standardized and highly reproducible qualitative method of antimicrobial susceptibility testing. It was developed in 1966, before broth microdilution by Kirby and Bauer, in response to the need for a more practical susceptibility test capable of measuring the susceptibility of multiple antibiotics simultaneously.20,25,28,29 Commercially prepared, filter paper disks containing a fixed concentration of an antibiotic are placed on solid media agar plates inoculated with a standardized inoculum of the infecting organism (1 to 2 × 108 CFU/mL). The plates are large enough to accommodate up to 12 different antibiotic disks at the same time (Figure 18-3).20 The plate is inverted to avoid moisture accumulation on the agar surface and incubated for 16 to 18 hours in ambient air at 35°C. During this incubation time, the antibiotic diffuses out of the disk into the surrounding media, with the highest concentration closest to the disk, as the bacteria multiply on the surface of the plates.20,25,29 The bacteria grow only in areas on the plate where the concentrations of the antibiotic are too low to inhibit bacterial growth. At the end of incubation period, the plates are examined for the inhibition of bacterial growth by measuring the diameter (in millimeters) of the clear zone of inhibition surrounding each filter paper disk. In general, the larger the zone size, the more active the antibiotic is against the organism.

FIGURE 18-3.
FIGURE 18-3.
A disk diffusion test where the diameters of all zones of inhibition are measured and those values are translated to categories of susceptible, intermediate, or resistant using the tables published by the CLSI. (Source: Reprinted with permission from Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:177–204.)

The diameter of the zone of inhibition has been correlated to the MIC of the antibiotic from broth or agar dilution against the infecting organism using regression analysis.20,25,28,29 CLSI has established interpretive criteria based on this relationship to categorize zone diameters as S, I, and R for each antibiotic against each organism.25,28,29 The results of disk diffusion test are considered qualitative because they only reveal the zone of inhibition and comparative activity of an antibiotic rather than an exact MIC.

The disk diffusion susceptibility test allows the simultaneous testing of several antibiotics in a relatively easy and inexpensive manner and provides flexibility in determining the antibiotics that will be tested for susceptibility, provided a filter paper disk for the desired antibiotic is available. However, the major disadvantages of disk diffusion include inability to generate an exact MIC and difficulty in determining the susceptibility of fastidious or slow-growing organisms.

Antibiotic Concentration Gradient Methods

Epsilometer test

The Epsilometer test, or Etest (bioMérieux) combines the benefits of broth microdilution with the ease of disk diffusion.11 The Etest method simultaneously evaluates the activity of numerous concentrations of an antibiotic using a single plastic strip impregnated on one side with a known, predefined concentration gradient of an antibiotic. One side of the Etest strip is marked with a numeric scale that depicts the concentration of antibiotic at that location on the reverse side of the test strip.9,20,25 Like disk diffusion, the Etest strip is applied onto a solid media agar plate that has been inoculated with a standardized concentration of the infecting bacteria. Several Etest strips can be placed on the same agar plate to provide the simultaneous susceptibility testing of several antibiotics.9,20,25 During overnight incubation, bacteria multiply on the agar plates as the antibiotic diffuses out of the Etest strip according to the concentration gradient. Bacterial growth occurs only in areas on the agar plate in which drug concentrations are below those required to inhibit growth. An elliptical zone of growth inhibition forms around the Etest strip where the MIC is read as the drug concentration where the ellipse intersects the plastic strip (Figure 18-4A and B).20,25

FIGURE 18-4.
FIGURE 18-4.
The Etest. (A) Individual Etest strips are placed on an inoculated agar surface. (B) After incubation, the MIC is read where the ellipse crosses the strip at the arrow. (Source: Reprinted with permission from Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:177–204.)

Etest results are reported as the exact MIC of the infecting bacteria with the corresponding CLSI susceptibility interpretation. The MIC results derived from the Etest correlate well with the results obtained using other susceptibility testing methods.9,20,25 The advantages of the Etest method include its ease of use and the ability to evaluate the susceptibility of several antibiotics simultaneously as well as the fact that the results yield an exact MIC, and the laboratory can choose the antibiotics to be tested. However, the Etest method is considerably more expensive than disk diffusion or broth microdilution methods, the results may be reader-dependent, and testing is limited to only those antibiotics for which an Etest strip is commercially available.

The Etest is currently used by some microbiology laboratories for the susceptibility testing of fastidious bacteria, such as S pneumoniae, H influenzae, and anaerobes, as well as for testing antibiotics in which a routine susceptibility test is not available (eg, antibiotic is not on standard broth microdilution panels used by the hospital) and when an exact MIC result is preferred.9,25

Specialized Susceptibility Tests

Additional tests may be performed in the microbiology laboratory to provide further information on the activity of an antibiotic against an organism. These specialty susceptibility tests may measure the bactericidal activity of the antibiotic (eg, MBC testing, time-kill curves, and serum bactericidal tests [SBTs]) or the activity of a combination of antibiotics against an infecting organism (eg, synergy testing using the checkerboard technique or time-kill studies). These tests are not routinely performed in most microbiology laboratories due to biological and technical difficulties, complexity in the interpretation of the results, and uncertain clinical applicability.22-24,31

Testing methods for determining bactericidal activity

Several methods measure the direct killing activity of an antibiotic against an organism and, if used, should be performed only for antibiotics that are generally considered to be bactericidal. As noted earlier, there are only a limited number of clinical circumstances in which this information may be useful. The determination of bactericidal activity may have the best clinical utility in the treatment of infections at anatomic sites where host defenses are minimal or absent, such as endocarditis, meningitis, and osteomyelitis, as well as in the treatment of severe and life-threatening infections in immunocompromised patients.9,20,22-24,31 Testing methods that determine the bactericidal activity of an antibiotic include the MBC test, time-kill assays, and SBTs.20,22-24,31

The MBC is the lowest concentration of an antibacterial agent that kills 99.9% of the bacterial inoculum, which represents a ≥3 log reduction in the original inoculum.11,22 The methodology for determination of the MBC has been previously described in detail in the section on broth macrodilution because it is an extension of that test. CLSI has developed guidelines to standardize the methodology for MBC testing.31 If the MBC is ≥32 times higher than the MIC or exceeds the achievable serum concentrations of the antibiotic, “tolerance” may be observed.20 Tolerance occurs when a normally bactericidal antibiotic only can inhibit the growth of bacteria based on MBC testing. MBC testing is not routinely performed by most laboratories because it is labor intensive with limited clinical use.20,22,31

Time-kill studies, also known as time-kill curves, measure the rate of bacterial killing over a specified period of time, which is in contrast to the MBC that measures the bactericidal activity at a single point in time following an incubation period.20,24,31 For time-kill studies, a standardized bacterial inoculum is placed into test tubes containing broth with several different concentrations of an antibiotic (usually the MIC and multiples of the MIC in separate tubes). Samples of the antibiotic-broth solutions are obtained at predetermined time intervals to evaluate the number of viable bacterial colonies present over the 24-hour incubation period.11,20,24,31 The number of viable bacteria present at each time point are plotted over time to determine the rate and extent of bacterial killing of the antibiotic against the organism. A ≥3 log reduction in viable bacterial counts is representative of bactericidal activity.11,20,24,31 Because it is labor and resource intensive, this test is not routinely performed in many clinical microbiology laboratories, but it is often used in the research setting.

The SBT, or Schlichter’s test, is similar to MIC and MBC testing, except the SBT measures the bacterial killing activity of the patient’s serum against their infecting organism after receiving a dose of an antibiotic.9,20,23,24,31-33 The methodology is similar to determining the MIC using broth macrodilution, but dilutions of the patient’s serum are used instead of 2-fold serial dilutions of an antibiotic.9,24,31-33 The patient’s serum is obtained at predefined intervals before and after a dose of an antibiotic, specifically at the time of expected peak concentration and at the time of expected trough concentration. The patient’s serum is then serially diluted and inoculated with a standardized concentration of the infecting organism. The SBT is the highest dilution of the patient’s serum that reduces the original standardized bacterial inoculum by ≥99.9%. The results of the SBT are reported as a titer, which represents the number of 2-fold serial dilutions of the patient’s serum that led to bacterial killing (eg, SBT = 1:16), with a higher titer indicating better activity against the organism.20,22,24,31-33 The CLSI has developed methodology standards for performance of the SBT.31,33 However, this test is not routinely performed by most microbiology laboratories because of technical difficulties. In addition, limited inconclusive data has been published regarding the clinical usefulness of SBTs in guiding therapy (only a few studies in the treatment of endocarditis, osteomyelitis, and serious infections in febrile neutropenia).22-24,31-33

Antimicrobial combination testing (synergy testing)

In the treatment of bacterial infections, there are several clinical situations in which combination antimicrobial therapy may be used. The decision to use combination therapy is primarily based on the severity of infection, the causative organism, and the type of infection. The potential benefits of combination antibiotic therapy include (1) expanding the antimicrobial spectrum of activity, especially empiric therapy for a life-threatening infection or for the treatment of polymicrobial infections; (2) producing synergistic bactericidal activity with the combination that is not observed with each agent alone, such as the use of ampicillin and gentamicin for the treatment of Enterococcal endocarditis; and (3) decreasing the emergence of resistant organisms, which has been observed in the treatment of tuberculosis (TB).24 Routine antimicrobial susceptibility tests measure the activity of an antibiotic against a particular organism. There are several tests, however, that evaluate the effects of combination antimicrobial therapy against an infecting organism (synergy testing), with the results being expressed as one of three types of activity11,20:

  1. Synergy: The activity of the antimicrobial agents in combination is significantly greater than the additive effects of each agent alone.

  2. Indifference: The activity of the antimicrobial agents in combination is similar to the additive effects of each agent alone.

  3. Antagonism: The activity of the antimicrobial agents in combination is less than the additive effects of each agent alone.

Therefore, before two antibiotics are used together, it may be useful to determine the effects of the combined antibiotics against the infecting organism, especially because some antibacterial combinations may produce suboptimal effects.

Synergy testing of an antimicrobial combination can be performed using the checkerboard technique, the time-kill curve technique, the disk diffusion assay, or the Etest method, with the checkerboard and time-kill curve techniques being most often used.20,24 The checkerboard technique is performed in macrodilution tubes or microdilution plates containing serial dilutions of the antibiotics alone and in combination. The tubes or plates are incubated with a standardized inoculum of the infecting bacteria for 24 hours. The effect of the antibiotic combination is determined by comparing the MICs of the agents when used in combination with the MICs of each agent alone. A synergistic combination displays lower MICs than when each agent is used alone. The time-kill curve method for combination therapy is similar to the time-kill curve method used to determine the rate of bacterial killing of a single agent, except that two antibiotics are added to the tubes in fixed concentrations. The effect of the antibiotic combination is determined by comparing the time-kill rates of combination therapy with the time-kill rates of each agent alone. A synergistic combination displays 100-fold or more killing activity than the most potent agent tested alone.11,20 In the clinical setting, synergy testing methods are not routinely performed due to their tedious, time-consuming methodologies, their expense, and their limited clinical applicability in predicting clinical outcome.11,20,24

Methods Detecting the Presence of Antibiotic Resistance Mechanisms

Detection of β-Lactamase Activity

To date, more than 2,700 different β-lactamase enzymes have been characterized.34 β-lactamase enzymes can be chromosomally-, plasmid-, or transposon-mediated and may be produced constitutively or inducibly. These enzymes cause hydrolysis of the cyclic amide bond in the β-lactam ring and, depending on the type of enzyme, may result in inactivation of one or numerous β-lactam antibiotics. It is important to understand the consequences of detecting a particular β-lactamase enzyme in an organism because certain enzymes produce resistance only to certain antimicrobials.20,23,34,35

Several methods detect the presence of β-lactamase enzyme depending on the organism and type of β-lactamase enzyme suspected. Some tests directly detect the presence of β-lactamase activity whereas others predict the presence of certain β-lactamase enzymes (such as the inducible AmpCs or the extended-spectrum β-lactamases [ESBLs]) based on resistance patterns and MICs derived from routine susceptibility tests.

The assays that directly detect β-lactamase activity include the acidimetric, iodometric, and chromogenic tests, which all measure the presence of β-lactamase enzyme by observing a color change based on reactions to different substrates.20,36 The chromogenic test is the most common test used by microbiology laboratories because of its reliability in detecting β-lactamase enzymes produced by many different bacteria.20,36 The chromogenic tests use chromogenic cephalosporins (nitrocefin, cefesone, or cefinase) incorporated into filter paper disks or strips that produce a color change when they are hydrolyzed by β-lactamase enzymes present in a clinical specimen once inoculated onto the disk or strip. Test tube assays using chromogenic cephalosporins can also be also used. A positive reaction using one of these direct β-lactamase tests for H influenzae, Moraxella catarrhalis, and N gonorrhoeae predicts resistance to only penicillin, ampicillin, and amoxicillin but not to other β-lactam antibiotics that are more stable to β-lactamase enzymes. A positive β-lactamase test for Staphylococcus spp. predicts resistance to penicillin, ampicillin, amoxicillin, carbenicillin, ticarcillin, and piperacillin.

Extended-spectrum β-lactamases are plasmid-encoded β-lactamase enzymes (eg, TEM, SHV, CTX-M) that hydrolyze most penicillins, cephalosporins (except the cephamycins and cefepime), and aztreonam.35 Historically, routine susceptibility tests using CLSI breakpoints did not reliably detect ESBL-producing organisms. Therefore, new CLSI interpretive criteria using lower MIC breakpoints for several cephalosporins and aztreonam for Enterobacterales were recently introduced to better detect resistance and obviate the need for ESBL screening and confirmatory tests (except for infection control or epidemiologic purposes). CLSI guidelines also outline criteria for performing screening and confirmatory tests for ESBLs that involve MIC and disk diffusion screening breakpoints for particular antibiotics using β-lactamase inhibitors.21,23,26-28,36,37 However, false-negative results may occur with these phenotypic ESBL confirmatory tests in the presence of an AmpC enzyme, which is not inhibited by clavulanic acid.23 Several automated susceptibility test systems, such as Vitek 2 and the Phoenix System, contain phenotypic ESBL detection tests that, when used with expert system software, are able to accurately detect ESBLs.37

AmpC β-lactamases are chromosomal- or plasmid-mediated β-lactamase enzymes that hydrolyze first-, second-, and third-generation cephalosporins and cephamycins, and also display resistance to some currently available β-lactamase-inhibitors, such as clavulanic acid, sulbactam, and tazobactam. Several gram-negative bacteria, such as S marcescens, P aeruginosa, indole-positive Proteus spp., Acinetobacter spp., Citrobacter freundii, and Enterobacter spp. (often referred to as the SPICE or SPACE bacteria) contain chromosomally-mediated, inducible AmpC enzymes that, when hyperproduced, can also hydrolyze penicillins and aztreonam in addition to the cephalosporins and cephamycins listed previously.37 AmpC hyperproduction can occur during the treatment of infection due to one of these organisms, especially when a strong inducer such as ceftazidime or clavulanic acid is used.35 Plasmid-mediated AmpC enzymes have also been reported in Klebsiella spp., Proteus mirabilis, Citrobacter koseri, and Salmonella spp., and often display an antibiotic susceptibility profile similar to chromosomally-mediated AmpC hyperproducers.37 All SPICE and SPACE bacteria should be assumed to be AmpC producers, so specific detection of AmpC production is not recommended.37 However, plasmid-mediated AmpC β-lactamases can be detected by demonstrating cephamycin hydrolysis using the AmpC disk test, the modified Hodge test (MHT), or the three-dimensional test.37

Carbapenemase enzymes have also emerged that may be chromosomal- (Stenotrophomonas maltophilia) or plasmid-mediated (eg, Pseudomonas aeruginosa, Acinetobacter spp., Klebsiella pneumoniae, etc.). Several plasmid-mediated carbapenemase enzymes have been characterized (KPC, VIM, OXA-4, NDM, and IMP) that hydrolyze carbapenems and most other β-lactam antibiotics, as well as display resistance to multiple other antibiotic classes.34,37 The modified Hodge test can be used for carbapenemase detection on isolates with elevated carbapenem MICs; however, it cannot differentiate between carbapenemase types.23,37

Because of the wide diversity of β-lactamase enzymes and the potential limitations of phenotypic ESBL-, AmpC-, and carbapenemase detection methods, several commercial tests have been developed to aid in the detection these enzymes, some of which include the RAPIDEC CARBA NP (bioMérieux, Durham, NC; phenotypic, colorimetric test that detects KPC, NDM, VIM, IMP, and OXA-48), Check-MDR assay (Wageningen, The Netherlands; microarray that detects TEM, SHV, CTX-M, some AmpCs, and NDM, VIM, IMP, OXA-48, and KPC), Expert CarbaR (Cepheid, Sunnyvale, CA; PCR that detects NDM, VIM, IMP, and OXA-48), FilmArray (bioMérieux; PCR that detects only KPC), and the Verigene (Luminex; PCR that detects CTX-M, KPC, NDM, VIM, OXA-48, and IMP).38

As mentioned earlier, the CLSI recently lowered the cephalosporin and carbapenem breakpoints for Enterobacterales in an attempt to better identify antibiotics with predictable efficacy against bacteria with multiple resistance mechanisms and eliminated the recommendation to perform specialized testing to detect ESBL-, AmpC-, or carbapenemase-mediated resistance. However, this recommendation has gained considerable criticism from many clinicians and microbiologists because detection of the exact mechanism of resistance is thought to be important for both treatment and epidemiologic purposes.37

High-Level Aminoglycoside Resistance

Aminoglycosides display relatively poor activity against Enterococcus spp. due to poor intracellular uptake (intrinsic, moderate-level resistance), so they should not be used alone in the treatment of infections due to enterococci. In addition, enterococci can acquire resistance to aminoglycosides through acquisition of genes that code for aminoglycoside-modifying enzymes (acquired resistance), which often leads to elevated aminoglycoside MICs (high-level aminoglycoside resistance or HLAR).23 Aminoglycosides (primarily gentamicin or streptomycin) may be considered with ampicillin, penicillin, or vancomycin to provide synergistic bactericidal activity, especially in the treatment of Enterococcal endocarditis or Enterococcal osteomyelitis. However, supplemental testing should be performed to detect the presence of HLAR, which predicts the lack of synergism between gentamicin or streptomycin and cell wall active agents against Enterococcus spp.20,23

The presence of HLAR can be evaluated using agar dilution (agar plates) or broth microdilution (wells) using high concentrations of gentamicin (500 mcg/mL) and streptomycin (2000 mcg/mL).23 The plates or wells are inoculated with a standardized suspension of the infecting Enterococcus spp. and incubated for 24 hours in ambient air.23 The growth of one or more Enterococcus spp. colonies on the agar plate or in the broth microdilution well demonstrates the presence of HLAR, signifying that the corresponding aminoglycoside cannot be used with a cell-wall active agent to achieve synergistic bactericidal activity. HLAR can also be detected using a disk diffusion method in which disks containing high concentrations of gentamicin (120 mcg) and streptomycin (300 mcg) are used.23 HLAR to gentamicin also confers resistance to tobramycin, netilmicin, and amikacin but not necessarily streptomycin, which should be tested independently.23 Testing for HLAR is usually performed only on Enterococcal isolates from infections that may require combination bactericidal activity, such as bacteremia, endocarditis, osteomyelitis, or meningitis.23

Tests for the Detection of MRSA, VISA, VRSA, and VRE

Several tests are available that can quickly detect or confirm the presence of methicillin-resistant S aureus (MRSA) or vancomycin-resistant enterococci (VRE). For the detection or confirmation of MRSA, the cefoxitin disk diffusion test, oxacillin–salt agar screening tests, culture-based chromogenic media, rapid latex agglutination (LA) tests, or molecular methods using real-time PCR can be used.23,30,39

The cefoxitin disk diffusion test is performed using routine CLSI procedures, with modified interpretive criteria used to detect mecA-mediated resistance in MRSA in which MRSA is reported for S aureus strains with a zone size of ≤21 mm.23,28 This test has also been useful in detecting methicillin-resistance in some coagulase-negative staphylococci.23 The oxacillin–salt agar screening tests have been widely used for the detection of mecA-mediated resistance MRSA, but they appear to lack sensitivity for the detection of strains that exhibit heteroresistance.23 A standard inoculum of S aureus is inoculated onto an agar plate containing Mueller-Hinton agar supplemented with 4% sodium chloride and 6 mcg/mL of oxacillin and incubated in ambient air at 33°C to 35°C for 24 hours.23 The growth of more than one colony indicates MRSA, which also confers resistance to nafcillin, oxacillin, cloxacillin, dicloxacillin, and all cephalosporins excluding ceftaroline. However, this test is not recommended for the detection of methicillin-resistance in other Staphylococcus spp.23

Selective chromogenic media are available to detect MRSA from surveillance specimens, all of which produce a characteristic pigment in the presence of MRSA with results available within 24 hours.23 There are also numerous rapid commercial LA tests to detect MRSA in clinical specimens by using highly specific monoclonal antibodies for the detection of penicillin-binding protein (PBP) 2a (also termed PBP 2′), the protein encoded by the mecA gene in MRSA.23,39

Several molecular tests for the detection of MRSA are commercially available that detect the mecA resistance determinant.38 Depending on the test, they may be used for surveillance testing (detecting colonization) or for the diagnosis of infection. Some examples of common molecular MRSA surveillance tests include the GeneOhm MRSA Assay (BD), the Xpert MRSA (Cepheid, Sunnyvale, CA), and the LightCycler MRSA Advanced Test (Roche Diagnostics, Indianapolis, IN), which all are FDA-approved PCR assays for the rapid, direct detection of nasal colonization by MRSA for the prevention and control of MRSA infection in healthcare institutions.23,30,38,39 These assays can detect the presence of MRSA directly from nasal swab specimens within 2 hours using real-time PCR that couples primers specific for mecA and the S aureus-specific gene orfX (sensitivity 93%, specificity 96%).39 Several PCR-based tests also exist for the diagnosis of infection due to MRSA and include the GeneOhm StaphSR Assay (BD; blood cultures), the XPert MRSA/SA BC and SSTI tests (Cepheid; blood cultures and skin/soft tissue infections), the Verigene Gram-Positive Blood Culture Nucleic Acid Test (Luminex), and the mecA XpressFISH (OpGen, Gaithersburg, MD), with results typically available within 1 to 2 hours of culture positivity.38

The CLSI reference broth microdilution method can accurately detect vancomycin intermediate S aureus (VISA, MIC 4 to 8 mcg/mL) and vancomycin-resistant S aureus (VRSA, MIC ≥16 mcg/mL) but may not consistently detect the presence of heteroresistant VISA.23 The use of brain heart infusion (BHI) agar plates with 6 mcg/mL of vancomycin (VRE screening plates described below) can be considered for the detection of S aureus strains with an MIC of ≥8 mcg/mL but is not useful for VISA strains with an MIC of 4 mcg/mL.23 Lastly, the disk diffusion test is unable to accurately detect VISA strains but detects VRSA strains mediated by vanA.23

Current automated susceptibility testing methods, including the Vitek 2 system and the BD Phoenix, are able to accurately detect the presence of VRE.23 VRE can also be detected using the vancomycin agar screen test, and is often performed on rectal swab specimens to detect carriers of VRE. A standard inoculum of the infecting Enterococcus spp. is inoculated onto an agar plate supplemented with BHI broth containing vancomycin 6 mcg/mL and incubated in ambient air for 24 hours.20,23 The presence of any growth demonstrates the presence of VRE. This test is most useful for detecting acquired vancomycin resistance (eg, vanA or vanB) in E faecalis and E. faecium, but it is not as useful for strains that display intrinsic resistance to vancomycin (eg, vanC), such as E. gallinarum and E. casseliflavus, in which MICs range from 8 to 16 mcg/mL (intermediate) and growth is variable on agar screening plates.

D-Zone Test for Detecting Inducible Clindamycin Resistance

Clindamycin resistance in staphylococci, S pneumoniae, and β-hemolytic streptococci is typically mediated by expression of the erm gene, which also confers resistance to macrolides, lincosamides, and streptogramin b (called MLSb-type resistance). MLSb resistance can be either constitutive or inducible, especially in staphylococci.20,23 Staphylococci (and β-hemolytic streptococci if susceptibility is performed) that are macrolide resistant but clindamycin susceptible should be evaluated for inducible clindamycin resistance using the d-zone test.20,23 The d-zone test is a disk diffusion procedure in which a 15-mcg erythromycin disk is placed 12 mm (streptococci) or 15 to 26 mm (staphylococci) apart from a 2-mcg clindamycin disk on an agar plate inoculated with the infecting organism.20,23 If inducible clindamycin resistance is present in the organism, the clindamycin zone of inhibition will be flattened on the side nearest the erythromycin disk, demonstrating the letter D in appearance. Organisms that display a flattening of the clindamycin zone are d-zone test positive and should be reported resistant to clindamycin in the final organism susceptibility report.

Special Considerations for Fastidious, Anaerobic, or Miscellaneous Bacteria

The susceptibility testing of fastidious bacteria (eg, H influenzae, N gonorrhoeae, and S pneumoniae) and anaerobes cannot be performed using standard broth microdilution, disk diffusion, or automated susceptibility testing methods because these organisms require more complex growth media and environmental conditions to support bacterial growth.36,40,41 The cultivation of fastidious bacteria or anaerobes may require media with supplemental nutrients, prolonged incubation times, and incubation in atmospheres with higher CO2 concentrations.36,40,41 Microbiology reference texts and CLSI standards have been developed to outline specific methodologies (broth dilution, disk diffusion, and automated methods), quality control guidelines, and interpretive breakpoint criteria that should be used for the susceptibility testing of these bacteria.21,25-29,36,40,41

The clinical significance of anaerobes as a cause of infection is more widely appreciated, and the susceptibility of anaerobes to various anti-infective agents is no longer predictable.9,40-43 The handling and processing of biologic specimens for anaerobic culture and susceptibility testing are extremely crucial to the validity of the results because most anaerobic bacteria of clinical importance are intolerant to oxygen.9,40 Specimens should be collected in appropriate anaerobic transport systems (commercially available vials or tubes) that contain specialized media and atmospheric conditions to support the growth of the anaerobic bacteria until the specimen is processed in the laboratory.40 Once collected, the specimens should be transported to the laboratory within minutes to hours of collection, processed for culture in anaerobic jars or chambers in the appropriate growth media, and incubated in anaerobic atmospheric conditions. The clinical specimens that provide the best yield for anaerobic culture include aspirated or tissue biopsy specimens.40

The identification of anaerobic bacteria by an individual hospital laboratory may be performed using one of three methods: (1) presumptive identification based on information from the primary growth plates, including the Gram stain results, patterns of growth on selective or differential media, plate and cell morphology, and results of various rapid spot and disk tests; (2) definitive identification based on the results of individual biochemical tests that detect the presence of preformed enzymes found in certain anaerobes; and (3) rapid identification of anaerobes using commercially available NA detection panels or MALDI-TOF.40,41 Many hospital laboratories do not have the resources for commercially available, anaerobic bacteria identification systems and rely on the first two methods for presumptive identification of anaerobic bacteria. If necessary, clinical isolates can be sent to a reference laboratory for further testing.

Most clinical microbiology laboratories do not currently offer routine susceptibility testing of anaerobic bacteria because of the uncommon occurrence of pure anaerobic infections, the uncertain role of anaerobes in mixed infections, the previous predictable susceptibility of anaerobic bacteria to antibiotics, the previous lack of standardization of antimicrobial susceptibility testing of anaerobes, and the technical difficulties in performing the tests.40-42 However, it is becoming apparent that routine antimicrobial susceptibility testing of anaerobic bacteria is necessary due to the increasing incidence of serious infections caused by anaerobic bacteria, the emerging resistance of anaerobic bacteria to multiple antibiotic agents, and the poor clinical outcomes observed when ineffective antibiotics are used for the treatment of infections caused by anaerobes.40-43

The susceptibility testing of anaerobic bacteria has undergone numerous methodological modifications and standardization over the past several years.41-43 The CLSI has recently published a standard outlining the clinical situations where anaerobic susceptibility testing should be considered, the methods of susceptibility testing that should be used, when and how surveillance susceptibility reporting should be performed, and the antibiotic agents that should be tested for susceptibility.42

Susceptibility testing for anaerobes should be performed in patients with serious or life-threatening infections such as endocarditis, brain abscess, osteomyelitis, joint infection, refractory or recurrent bacteremia, and infection of prosthetic devices or vascular grafts.40-42 Susceptibility testing should also be performed in patients with persistent or recurring anaerobic infections despite appropriate antibiotic therapy.41,42 Lastly, susceptibility testing of anaerobic bacteria should be periodically performed within geographic areas or individual institutions to monitor regional susceptibility patterns of anaerobic bacteria over time.40-42

The recommended anaerobic susceptibility testing methods include agar dilution and broth microdilution using supplemented Brucella broth, both of which can be reliably performed by most clinical microbiology laboratories.40-43 The agar dilution method is the gold standard reference method that can be used for susceptibility testing of any anaerobic bacteria, whereas the broth microdilution method has been validated only for antimicrobial susceptibility testing of Bacteroides spp. and Parabacteroides spp.41,42 In contrast to agar dilution, the broth microdilution method can evaluate the susceptibility of multiple antibiotics simultaneously, and several microdilution panels are now commercially available for routine susceptibility testing, including Anaerobe Sensititre panel (ANO2, Thermo Fisher Scientific) and Oxoid ANA MIC Panel (Thermo Fisher Scientific). The general methodology for each of these tests is similar to those described previously for aerobic bacteria. Etest strips can also be used for anaerobe susceptibility testing, and results appear to correlate well with agar dilution. In addition, β-lactamase testing of anaerobes can be performed according to CLSI guidelines using chromogenic disks.41-43

Because routine antimicrobial susceptibility of anaerobes is not performed by all hospital microbiology laboratories or for all anaerobic isolates, antibiotic therapy for infections caused by anaerobes is usually selected empirically based on susceptibility reports published by reference laboratories.41,42 However, if susceptibility testing is performed on an individual anaerobic isolate, the results should be used to guide the anti-infective therapy for the patient.

Lastly, several miscellaneous (often uncommon) pathogenic bacteria are difficult to detect or cultivate using the standard microbiologic procedures outlined previously. These organisms often pose a diagnostic dilemma because they often require specialized testing for identification. It is beyond the scope of this chapter to describe all specialized testing methods that are available to detect these organisms; however, an abbreviated list can be found in Table 18-10.4,13,44-61

TABLE 18-10.
Laboratory Tests for the Detection of Miscellaneous Organisms

ORGANISM

TYPE OF ORGANISM

CLINICAL FINDINGS AND INFECTIONS

DIAGNOSTIC METHOD

POSITIVE RESULT

REFERENCE

Bordetella pertussis (Whooping cough)

Bacteria

Upper respiratory tract symptoms, characteristic whooping cough, pneumonia

Culture

Growth within 3–7 days; more sensitive when performed early in course of infection

44

DFA, ELISA

Rapid detection of B. pertussis antibodies; should be used in conjunction with culture due to variable specificity

PCR

Direct detection in 1–2 days; most sensitive when performed early in course of infection

Borrelia burgdorferi (Lyme disease)

Spirochete

Erythema migrans, pericarditis, arthritis, neurologic disease

Culture

Not routinely performed due to difficulty and low sensitivity; long incubation (hold cultures for up to 12 wk)

13,45,46

ELISA (or rarely IFA); first step screening test

Measures IgM (appears within 1–2 wk) and IgG (appears within 4–6 wk) antibodies against B burgdorferi

WB if ELISA borderline or positive, second step confirmatory test

Measures IgM and IgG antibodies against B burgdorferi

PCR (confirm)

Can detect low numbers of spirochetes; especially useful in synovial fluid

Brucella spp.

Bacteria

Systemic infection (can involve any organ); spondylitis, arthritis, endocarditis

Culture

Growth within 7 days, but cultures should be held for 3 wk

47

SAT, MAT

Detects antibodies to most Brucella spp.; titer of ≥1:160 is diagnostic in conjunction with appropriate clinical scenario

ELISA

Useful for detection of chronic or past brucellosis; most useful for diagnosis of neurobrucellosis

PCR

Detection of Brucella-specific DNA sequences; not routinely available in most laboratories

Chlamydia pneumoniae

Atypical bacteria

Upper respiratory tract infections, pharyngitis; pneumonia

Culture

Monoclonal antibodies detect organism in culture

4,13,48

CF

4-fold rise in antibody titer between paired sera (acute and convalescent samples)

MIF

4-fold rise in antibody titer between paired sera (acute and convalescent samples) or a single serum sample with an IgM titer of 1:16 or an IgG titer 1:512

PCR

Detection of C pneumoniae DNA

Clostridioides difficile (Pseudomem-branous colitis)

Anaerobic bacteria

Pseudomembranous colitis, diarrhea

Toxigenic culture using CCFA growth media

Growth within 48 hr; most sensitive test

49

CCNA or EIA toxin test

Detection of toxin A or B activity; cell cytotoxicity test more sensitive than EIA

PCR

Rapid sensitive and specific detection of Clostridioides difficile tcdB gene

Glutamate dehydrogenase (GDH) assays

Detects GDH, an enzyme present in all Clostridioides difficile isolates; cannot differentiate between toxigenic and nontoxigenic strains

Coxiella burnetti (Q fever)

Bacteria

Acute or chronic systemic illness, pneumonia, hepatitis, endocarditis

Culture with DFA

Growth in 6–14 days, organism detected by DFA

13,48

IFA, EIA, CF

IgM titer of 1:50 or an IgG titer of 1:200

Cryptosporidium parvum

Protozoa

Acute diarrhea (self-limiting to severe), abdominal pain, dehydration

Modified acid fast, Ziehl-Neelsen or Kinyoun staining

Detection of oocysts in stool or intestinal scrapings

50

DFA using a monoclonal antibody against oocyst

Detection of oocysts in stool or intestinal scrapings

EIA

Detection of C. parvum antigen in stool or intestinal scrapings

PCR

Detection and differentiation of Cryptosporidium spp.

Ehrlichia spp.

Bacteria

HME - fever, myalgia, headache, malaise, rash (more common in children), nausea, vomiting, diarrhea, leukopenia, thrombocytopenia, elevated hepatic transaminases; may be life-threatening

IFA serology

4-fold rise in antibody titer between paired sera (acute and convalescent samples)

48,51,52

Peripheral blood smear Wright-Giemsa or Diff-Quik stain

Detect morulae (cytoplasmic vacuoles)

PCR

Detection of E chaffeensis or E phagocytophilum DNA sequences

Entamoeba histolytica

Protozoa

Amebiasis: intestinal (colitis, diarrhea) and extraintestinal (liver abscess)

Stool exam for ova and parasites

Detection of trophozoites and cysts

50

Serology (IFA)

Detection of E histolytica antibodies with titer ≥1:200

Antigen detection on fresh stool samples

Detection of E histolytica or E dispar specific antigen

PCR

Detection and differentiation of E histolytica or E dispar

Giardia spp.

Protozoa

Acute diarrhea (self-limiting to severe), malabsorption syndromes, low-grade fever, chills, abdominal pain

Stool exam for ova and parasites

Detection of trophozoites and/or cysts

50

Wet preps or stains of duodenal material

Detection of trophozoites and/or cysts

EIA, DFA, or ICA antigen detection assays

Detection of trophozoite and/or cyst antigens

Helicobacter pylori

Bacteria

Peptic ulcer disease

Urea breath test

Positive test indicative of the presence of organism

53

LA serologic tests

Detect IgG antibodies against H pylori

Stool antigen tests using monoclonal antibodies (ELISA)

Detection of H pylori antigen

Urease test on antral biopsy specimen

Positive test indicative of active infection

PCR

Detection of H pylori DNA

Legionella pneumophila

Atypical bacteria

Pneumonia

Culture using specialized media

Growth in 3–5 days

4,54

DFA staining

Binds to L. pneumophila antigen to produce fluorescence

IFA serology

4-fold rise in antibody titer between paired sera (acute and convalescent samples)

Urinary antigen detection (EIA, RIA, ICA)

Detects Legionella pneumophila serogroup 1 antigen only

PCR

Detection of Legionella spp. DNA

Leishmania spp.

Protozoa

Cutaneous, mucocutaneous, or visceral (VL, kala-azar) infection; can infect reticuloendothelial system

Giemsa staining and light microscopy

Amastigotes within the specimen

55

Culture

Growth of promastigotes

IFA, DAT, ELISA (VL)

Detection of antileishmanial antibodies in blood or serum

LA (VL)

Detection of leishmanial antigen in urine

PCR (VL)

Detection of Leishmania DNA

Leptospira spp.

Spirochetes

Leptospirosis (self-limiting with fevers, chills, myalgia, headache, aseptic meningitis); icteric leptospirosis (severe form associated with jaundice, bleeding, and renal failure)

Dark-field microscopy or immunofluorescence

Detection of motile leptospires

13,46

Culture

Growth within 6 wk

Serology using MAT

4-fold or greater rise in agglutinating antibody titer between paired sera (acute and convalescent samples)

ELISA

Detection of leptospiral antibodies

PCR

Detection of leptospiral DNA

Mycoplasma hominis

Atypical bacteria

Urogenital tract infections, including prostatitis, PID, bacterial vaginosis, urethritis; systemic infection in neonates and immunocompromised patients

Culture using selective media

Growth within 5 days

13,56

Mycoplasma pneumoniae

Atypical bacteria

Pneumonia, tracheobronchitis, pharyngitis

EIA, IFA, CF serology

4-fold or greater rise in antibody titer between paired sera (acute and convalescent samples)

4,13,56

NAAT, PCR

Detection of M pneumoniae DNA

Plasmodium falciparum, P. vivax, P. ovale, P. malariae (Malaria)

Protozoa

Symptoms include high fever (cyclic with P. vivax, P. ovale, P. malariae), chills, nausea, vomiting, severe headache, anemia, abdominal pain; life-threatening with P. falciparum

Thick and thin blood films stained with Giemsa or Wright’s stain (gold standard)

Presence of malarial parasites

55

Fluorescent-assisted microscopy with acridine orange

Detection of fluorescence when dyes are taken up by the nucleus of the parasite

ICA antigen assays

Detection of malaria-specific antigens

PCR

Detection of malaria-specific DNA sequences

Pneumocystis jirovecii (carinii; PCP)

Fungus with protozoal characteristics

Pneumonia, extrapulmonary infection

Microscopic exam after Giemsa or methenamine silver stain of induced sputum, BAL specimen, or tissue biopsy

Detection of trophic or cystic forms

57

DFA or IFA

Detection of cysts or trophozoites

PCR

Detection of P jirovecii-specific DNA

1,3-β-d-glucan serum test

Positive values >100 pg/mL

Rickettsia rickettsii (Rocky Mountain spotted fever)

Rickettsia

Fever, chills, headache and rash in patient with recent tick bite; myalgias, malaise, nausea, vomiting, abdominal pain, focal neurologic findings; small vessel vasculitis may result in life-threatening complications

IFA (gold standard)

4-fold or greater rise in IgM or IgG antibody titers between paired sera (acute and convalescent samples)

48,52

EIA or LA

Detection of IgM or IgG antibodies

PCR

Detection of R rickettsii DNA sequences

Strongyloides stercoralis

Parasite

Abdominal infection; disseminated infection (hyperinfection syndrome with pneumonitis, sepsis)

Stool exam for ova and parasites

Detection of adult worms, eggs, and/or larvae

58

Taenia solium

Tapeworm

Neurocysticercosis (infection within brain tissue) causing seizures, headache, focal neurologic deficits; muscular and subcutaneous abscesses

Serology by EITB

Detection of antibodies to T solium glycoprotein antigens

59

ELISA on CSF

Detection of anticysticercal antibodies or cysticercal antigens

Toxoplasma gondii (Toxoplasmosis)

Protozoa

Encephalitis, myocarditis, lymphadenitis, polymyositis, chorioretinitis, toxoplasmosis during pregnancy, congenital toxoplasmosis

Serology testing

Positive IgG antibody

60,61

Giemsa or Diff-Quik stain of CSF or body fluid/tissue

Demonstration of tachyzoites

PCR

Detection of T gondii-specific DNA

Ureaplasma urealyticum

Atypical bacteria

Urogenital tract infections, including prostatitis, PID, bacterial vaginosis, urethritis; systemic infection in neonates and immunocompromised

Culture using selective media

Growth within 5 days

13,56

PCR

Detection of NA or gene targets

BAL = bronchoalveolar lavage; CCFA = cycloserine cefoxitin fructose agar; CCNA = cell cytotoxicity neutralization assay; CF = complement fixation; DAT = direct agglutination test; EITB = enzyme-linked immunoelectrotransfer blot; GDH = glutamate dehydrogenase; HME = human monocytic ehrlichiosis; ICA = immunochromatographic assay; IFA = immunofluorescent assay/indirect fluorescent antibody; MAT = microagglutination test; MIF = microimmunofluorescence; NAAT = nucleic acid amplification test; O&P = ova and parasites; PID = pelvic inflammatory disease; RIA = radioimmunoassay; SAT = serum agglutination test; VL = visceral leishmaniasis; WB = western blot.

Source: References 4,13,4461.

Methods for Reporting Susceptibility Results

Individual Isolate Susceptibility Reports

When a bacterial isolate is recovered from a clinical specimen, the identification and susceptibility results are compiled in a report that is available electronically or via a hard copy in the patient’s chart. The bacterial identification and antibiotic susceptibility report often contains the following information: the patient’s name, medical record number, the date and time of specimen collection, the source of specimen (eg, blood, wound, urine), the bacteria that were identified (if any), and the list of antibiotics tested for susceptibility with the MIC or disk diffusion result and CLSI interpretive category, as shown in Figure 18-5.20,62 Additionally, the site of blood culture collection (eg, drawn peripherally versus central line) and the time to positivity are occasionally reported. In some hospitals, the individual isolate susceptibility report may also contain information regarding the usual daily dose and cost of antibiotics.

FIGURE 18-5.
FIGURE 18-5.
Example of microbiology laboratory report with bacterial identification and antibiotic susceptibility.

Once bacterial culture and susceptibility results are available, this information should be used to change the patient’s empiric antibiotic regimen, which usually covers a broad spectrum of bacteria, to a more directed antibiotic regimen targeting the infecting bacteria and antibiotic susceptibility (de-escalation). The directed antibiotic regimen should be chosen based on clinical and economic factors, some of which include the severity of infection, the site of infection, the activity (MIC value) of the antibiotic against the infecting organism, the proven efficacy of the antibiotic in the treatment of the particular infection, the overall spectrum of activity of the antibiotic (a narrow spectrum agent is preferred), the end-organ function of the patient, the presence of drug allergies, the route of administration (oral versus parenteral), the daily cost of the antibiotic, and so on. The susceptibility report provides some of the information necessary for the de-escalation of antibiotic therapy, namely, the site of infection, the infecting organism(s), and the susceptibility of the infecting organism(s).

As seen in the sample susceptibility report in Figure 18-5, several antibiotics may display activity against the infecting bacteria, often with different MICs. It is not always advantageous to choose the antibiotic with the lowest MIC against a particular organism on a susceptibility report. As discussed earlier in this chapter, antibiotics have different MIC breakpoints corresponding to S, I, and R (and SDD for select antibiotics) for each bacterium based on several factors. Some drugs, such as piperacillin–tazobactam, are assigned higher MIC breakpoint values for susceptibility because they achieve higher serum and site concentrations than other antibiotics. Because of this, a simple number comparison of the MIC between antibiotics should not be performed. The choice of antibiotic should be based on the knowledge of the MICs that are acceptable for a particular antibiotic–bacteria combination, the site of infection, the penetration of the antibiotic to the site of infection, as well as the clinical and economic parameters listed previously. In the sample report in Figure 18-5, oxacillin (nafcillin) or cefazolin would be an acceptable choice for the treatment of S aureus bacteremia in a patient without drug allergies because these agents are active against the infecting organism, have been demonstrated to be effective in the treatment of systemic staphylococcal infections, have a relatively narrow spectrum, and are inexpensive. Minicase 2 is an example illustrating the use of a bacterial culture and susceptibility report in the antibiotic decision-making process.

The decision regarding the specific antibiotics that will be routinely reported in an individual susceptibility report for a bacterial isolate is typically based on input from a hospital or institutional multidisciplinary committee (eg, Antimicrobial Subcommittee, Infectious Diseases Subcommittee, Antimicrobial Stewardship Team) comprised of infectious diseases physicians, infectious diseases pharmacists, and representatives from the Infection Control Committee and the microbiology laboratory. The choice of specific drugs to report is often based on the hospital formulary, the level of control of antibiotic use that is desired, and the tests that are used by the microbiology laboratory for susceptibility testing. Tables that outline the antibiotics that should be routinely tested and reported for each bacterium can be found in the CLSI Performance Standards and Guidelines for Antimicrobial Susceptibility Testing.21,26-28

Several methods can be used for reporting antibiotic susceptibility of individual bacterial isolates, including general reporting, selective reporting, and cascade reporting. General reporting involves reporting all antibiotics that were tested for susceptibility against the organism without any restrictions. Selective reporting involves reporting certain antibiotic susceptibility test results on an individual bacterial isolate based on defined criteria, such as the bacteria identified, the site of infection, antibiotics available on the hospital formulary, etc. An example of selective reporting would be the exclusion of cefazolin from the susceptibility report of a CSF sample growing E coli because cefazolin is not a suitable treatment option for meningitis. Cascade reporting involves the preferential release of susceptibility information for first-line choices for the treatment of a particular organism or infection (usually narrow spectrum and inexpensive), with the reporting of second-line antibiotic susceptibility results (usually broad spectrum and costly) only if the first-line agents are inactive against the infecting organism or are inappropriate for the treatment of the particular infection. An example of cascade reporting is the reporting of amikacin susceptibility results against P aeruginosa only if the organism displays resistance to both gentamicin and tobramycin, which are less expensive aminoglycoside agents. Both selective reporting and cascade reporting are often used as antimicrobial stewardship activities to control the inappropriate use of broad-spectrum or expensive antibiotics.62

Hospital Susceptibility Reports (Hospital Cumulative Antibiograms)

Most hospitals prepare and publish an annual cumulative report of antimicrobial susceptibility profiles of the bacteria that have been isolated from patients within their hospital, healthcare system, or institution, called a cumulative antibiogram. For the cumulative antibiogram to be clinically useful, the susceptibility data from patient isolates should be appropriately collected, analyzed, and reported according to the CLSI guidelines, which are outlined in Table 18-11.62

TABLE 18-11.
CLSI Recommendations for Cumulative Antibiogram Development
  1. To serve as a continuously useful tool to guide appropriate empiric antibiotic therapy, the cumulative antibiogram should be compiled, analyzed, and reported at least annually.

  2. Only the first clinical isolate of a given species of bacteria per patient per analysis period (annually if that is the time frame of the antibiogram) should be included in the cumulative susceptibility report regardless of site of isolation, susceptibility pattern of the bacteria, or other phenotypic characteristics. The inclusion of duplicate clinical isolates from the same patient will lead to overreporting of bacterial resistance.

  3. To provide a reasonable statistical estimate of susceptibility, only species of bacteria in which at least 30 isolates have been collected, tested, and reported during the time period of the antibiogram should be included.

  4. Data from isolates recovered during surveillance cultures (eg, MRSA and VRE), environmental cultures, or other nonpatient sources should not be included in the antibiogram.

  5. The cumulative susceptibility report should include all antibiotics that were tested for susceptibility, regardless of whether they were reported in the final susceptibility report for the individual patient.

  6. Only bacterial isolates for which all routine antibiotics have been tested for susceptibility should be included. Results of agents selectively or supplementally tested should not be included in the cumulative susceptibility report. For example, if only isolates resistant to primary agents were then analyzed for susceptibility to secondary agents, this will bias the resistance results toward higher levels of resistance to the secondary agents.

  7. Data may be stratified and reported by age, unit in which the isolate was collected (eg, MICU, SICU, outpatient clinic), or anatomic site of collection (eg, blood isolates, CSF isolates, and urine isolates) as long as duplicate patient isolates are removed and there are a sufficient number of isolates collected (> 30) during the time frame of the antibiogram.

MICU = medical intensive care unit; SICU = surgical intensive care unit.

Source: Reference 62.

The cumulative antibiogram contains information on the percent of isolated bacteria that were susceptible to antibiotics tested over the time frame of the antibiogram, as illustrated in Figure 18-6.62 This percent susceptibility information is derived by dividing the number of organisms susceptible to a particular antibiotic by the total number of single-patient isolates collected and reported (with duplicate patient isolates removed). The calculations can be performed either manually or through the use of automated systems that have been programmed using appropriate definitions to remove duplicate patient isolates. Cumulative antibiograms usually contains separate data tables for the susceptibility reporting of gram-positive, gram-negative, and anaerobic bacteria (if performed).

FIGURE 18-6.
FIGURE 18-6.
Example of a hospital antibiogram report.

The specific data published in the cumulative antibiogram should be based on input from the hospital/healthcare system’s multidisciplinary committee (eg, Infectious Diseases Subcommittee, Antimicrobial Stewardship Committee) that is often comprised of infectious diseases physicians, infectious disease pharmacists, infection preventionists, and the microbiology laboratory. The cumulative antibiogram report typically contains information on the antibiotic susceptibility patterns of isolates obtained from patients in the hospital (either admitted with infection or who developed infection in the hospital) but may also include the susceptibility of organisms causing infection in outpatients if the hospital/medical center also serves a substantial outpatient population. In addition, each hospital or healthcare system may further stratify their susceptibility data by various parameters, such as patient care unit (eg, Burn Unit, MICU, Pediatric Unit, Med-Surg Unit, outpatient clinic, nursing home), patient age, site of infection (eg, bloodstream isolates, UTI isolates), patient characteristics (eg, cystic fibrosis, transplant patients, hematology/oncology patients), or by organism (eg, susceptibility of S aureus). For some organisms, the cumulative antibiogram only will contain information regarding the presence of bacterial resistance mechanisms, particularly when routine susceptibility testing is difficult to perform, such as in the case of H influenzae in which the percentage of isolates that produce β-lactamase enzyme during the time period of the cumulative antibiogram will be reported. Other information that may be incorporated into a cumulative antibiogram includes graphs demonstrating resistance trends, antibiotic dosing guidelines, recommended empiric antibiotic choices based on infection type, and antibiotic cost data.62

Using Laboratory Test Results to Guide the Choice of a Directed Antibiotic Regimen for the Treatment of Pyelonephritis

Diana J., a 27-year-old woman, presents to the urgent care clinic with reports of urinary frequency and urgency, pain on urination, and hematuria for the past 2 days. She also reports fever of 101.6°F and intractable nausea and vomiting during the past 24 hours. At presentation, she is febrile (102.3°F), hypotensive (90/60 mm Hg), and lethargic; physical exam reveals right costovertebral angle and suprapubic tenderness. A urine dipstick performed in the clinic is leukocyte esterase positive, and a urine pregnancy test is negative. Because she appears ill, the clinic physician sends the patient to the local emergency department (ED) for admission. Her past medical history is significant for recurrent UTIs, with three episodes over the past 6 months that have required antibiotic therapy, including oral trimethoprim–sulfamethoxazole and oral ciprofloxacin. She reports no known drug allergies. Upon admission to the ED, urinalysis, urine culture, and blood cultures are performed.

QUESTION: What is an appropriate recommendation for emipric antibiotic therapy for this patient?

DISCUSSION: Based on her presenting symptoms and the findings on her physical examination, this patient most likely has acute pyelonephritis, an upper tract UTI, making the acquisition of urinalysis, urine culture, and blood cultures important for guiding directed antimicrobial treatment. This is especially important in this patient because she has received multiple recent courses of antibiotics for her past UTIs, which put her at risk for infection with resistant bacteria. Because she is hypotensive and is experiencing significant nausea and vomiting, she should initially receive empiric parenteral antibiotic therapy that displays activity against the likely causative organisms of pyelonephritis and has proven efficacy in the treatment of complicated UTIs (eg, ceftriaxone because she does not have any antibiotic allergies). The patient should receive ceftriaxone as empiric therapy, which can be de-escalated to cefazolin based on the results of the blood and urine culture and susceptibility test results that follow.

URINALYSIS: Yellow, cloudy; pH 7; specific gravity 1.015; protein negative; RBC trace; WBC 50 to 100/hpf; leukocyte esterase positive; nitrite positive.

Midstream Urine Culture/Susceptibility: > 100,000 CFU/mL of E coli

ANTIBIOTIC TESTED

MIC RESULT

CLSI INTERPRETATION

Ampicillin

>32 mcg/mL

R

Ampicillin–sulbactam

8 mcg/mL

S

Cefazolin

1 mcg/mL

S

Ceftriaxone

1 mcg/mL

S

Imipenem

1 mcg/mL

S

Gentamicin

0.5 mcg/mL

S

Ciprofloxacin

4 mcg/mL

R

Trimethoprim–sulfamethoxazole

>4/76 mcg/mL

R

R = resistant; S = susceptible.

Blood Culture/Susceptibility: E coli

ANTIBIOTIC TESTED

MIC RESULT

CLSI INTERPRETATION

Ampicillin

>32 mcg/mL

R

Ampicillin–sulbactam

8 mcg/mL

S

Cefazolin

1 mcg/mL

S

Ceftriaxone

1 mcg/mL

S

Imipenem

1 mcg/mL

S

Gentamicin

0.5 mcg/mL

S

Ciprofloxacin

4 mcg/mL

R

Trimethoprim–sulfamethoxazole

>4/76 mcg/mL

R

R = resistant; S = susceptible.

The cumulative antibiogram is a useful tool for selecting empiric antibiotic therapy, where an antibiotic is selected based on the local susceptibility patterns of the most likely infecting organism causing the patient’s infection (Table 18-7) while waiting for the results of culture and susceptibility tests, as described in Minicase 3.62 Antibiotic therapy must often be initiated at the suspicion of infection because many infectious diseases are often acute where a delay in treatment may result in significant morbidity or mortality (eg, meningitis and pneumonia). Once the culture and susceptibility results of the infecting bacteria are known, antibiotic therapy should be deescalated or directed, if necessary, to an agent with more targeted activity against the organism

Using the Cumulative Antibiogram to Choose Empiric Antibiotic Therapy

David M. is a 45-year-old man who sustained multiple traumatic injuries after a motorcycle accident. He has required multiple surgeries over the past 10 days for fracture stabilization. In the last 12 hours, he has spiked a temperature to 39°C and has developed shaking chills. His other vital signs are stable, and his physical exam does not demonstrate any significant focal findings. Urinalysis, urine culture, and blood cultures are performed to determine the potential etiology for his new fever. In addition, a chest radiograph is obtained, which does not demonstrate any pulmonary infiltrates. The microbiology laboratory calls the surgical floor later that day to report that the blood culture results are positive for gram-negative rods. The patient is allergic to penicillin (nonurticarial rash); the local hospital antibiogram is pictured in Figure 18-6.

QUESTION: What empiric antibiotic therapy should be used to treat this patient’s gram-negative rod bacteremia while waiting for the culture and susceptibility results?

DISCUSSION: Nosocomial gram-negative bacteremia is a potentially life-threatening infection that requires early, aggressive antibiotic therapy. The choice of whether to use monotherapy or combination therapy while waiting for culture and susceptibility results in this setting often depends on the clinical condition of the patient and local susceptibility patterns. Combination antibiotic therapy might be considered initially if the patient is critically ill (septic shock) from the bacteremia because it might provide some synergistic antibacterial activity as well as enhanced coverage against a wide range of potential infecting bacteria. Based on the hospital antibiogram in Figure 18-6, it is desirable to choose an antibiotic that displays good activity (>85% susceptible) against gram-negative bacteria isolated at the institution (eg, P aeruginosa, E coli, K pneumoniae, S marcescens, and Enterobacter cloacae) and choose an antibiotic with proven efficacy in the treatment of bacteremia. Because the patient is clinically stable and displays only a rash to penicillin, some useful therapeutic options based on review of the hospital antibiogram include meropenem, ceftazidime, or cefepime. If the patient were to clinically deteriorate on monotherapy, an aminoglycoside (tobramycin) or a fluoroquinolone (ciprofloxacin) could be added to the carbapenem or cephalosporin while waiting for the culture and susceptibility results. The antibiotic regimen could then be modified to more directed therapy, if possible, once the final culture and susceptibility results were available.

Surveillance Susceptibility Testing of Large Numbers of Isolates

Surveillance susceptibility testing is a useful method to monitor the susceptibility of bacteria to antimicrobial agents over time and can be performed in an individual hospital or within a geographic location (eg, regionally, nationally, and internationally).20 Surveillance studies typically report the overall susceptibility of the bacteria to particular antibiotics using CLSI breakpoints, along with other susceptibility parameters, such as the MIC50 and the MIC90. To determine the MIC50 or MIC90, the MIC values from the bacterial population studied are arranged in ascending order, where the MIC50 is the MIC value representing 50% of the bacterial population (the MIC value of the isolate that represents 50% of the bacterial population studied) and the MIC90 is the MIC that represents 90% of the bacterial population (the MIC value of the isolate that represents 90% of the bacterial population studied). The MIC90 value is usually higher than the MIC50 value. This information is useful for detecting the emergence of subclinical antibiotic resistance in which the MIC50 and MIC90 of a particular agent may be increasing over time but are still below the MIC susceptibility breakpoint.

Additional Considerations When Interpreting Susceptibility Results

The successful treatment of a patient’s infection involves an understanding of the interactions among the patient, the infecting organism, and the antibiotic. It is important to note that antimicrobial susceptibility testing only measures one of these factors, namely, the activity of the antibiotic against the infecting organism in a laboratory setting. The current methodologies for antibiotic susceptibility testing are unable to reproduce the interaction between the antibiotic and the bacteria at the site of infection in which a multitude of host factors (eg, immune system function, concomitant disease states) and drug factors (eg, pharmacokinetic parameters, including concentration of free drug at the site of infection and protein binding) play an integral role.

LABORATORY TESTS USED FOR DIAGNOSIS OF SPECIFIC INFECTIONS

Bacterial Meningitis

Meningitis is an infectious diseases medical emergency requiring prompt, accurate diagnosis and treatment. Meningitis may be caused by bacteria, viruses, fungi, or mycobacteria, and it produces a resulting clinical presentation of acute or chronic meningitis depending on the causative organism. In a patient with suspected meningitis, a lumbar puncture is performed to obtain CSF for laboratory analysis to aid in the diagnosis of the infection, including the potential causative organism.2,63-67 In patients who present with papilledema, altered consciousness, new-onset seizures, or focal neurologic findings, a head computed tomography may be performed prior to the lumbar puncture to exclude the presence of a space-occupying lesion, which may put the patient at risk for brain herniation after lumbar puncture.63,66 A lumbar puncture involves the aseptic insertion of a spinal needle into the subarachnoid space at the lumbar spine level (between L3, L4, or L5) for the aspiration of 5 to 20 mL of CSF for analysis.63,64 When inserting the spinal needle, the opening pressure may be measured (normal opening pressure is 50 to 195 mm H20 in adults) and is often elevated in patients with meningitis (especially C neoformans meningitis) and concomitant cerebral edema or intracranial focus of infection.63,66 The CSF should be placed in three to four separate sterile screw-cap tubes and immediately transported to the laboratory for rapid processing. The first two tubes of CSF are used for microbiologic (eg, Gram stain, fungal stains, acid-fast bacilli stain, culture, and antigen detection) and chemical studies (eg, general appearance, glucose, and protein), whereas the last tubes are used for determination of the WBC count and differential. The typical CSF chemistry, hematology, and microbiologic findings in patients with meningitis caused by different pathogens are listed in Table 18-12.63-67

TABLE 18-12.

Typical CSF Findings in Patients with Meningitis

NORMAL

BACTERIAL MENINGITIS

VIRAL INFECTION

FUNGAL MENINGITIS

TUBERCULOUS MENINGITIS

Opening pressure (mm H20)

<180

>195

WBCs (count/mm3)

0–5

<30 (newborns)

1,000–20,000 (mean 800)

50–2,000 (mean 80)

20–2,000 (mean 100)

5–2,000 (mean 200)

WBC differential

No predominance

≥80% PMNs

>50% lymphs, 20% PMNs

>50% lymphs

>80% lymphs

Protein (mg/dL)

<50

>100

30–150

40–150

>50

Glucose (mg/dL)

(CSF/blood glucose ratio)

45–100

(two-thirds of serum)

<45

(< one-half of serum)

45–70

30–70

<40

Gram stain (% positive)

60–90

Negative

Negative

37–87 (AFB smear)

PMNs = polymorphonuclear leukocytes; lymphs = lymphocytes.

Source: References 6367.

Chemistry and Hematology

In patients with meningitis, the CSF often appears cloudy because of the presence of WBCs, protein, and bacteria.66 The chemistry and hematology results from the CSF analysis directly correlate with the probability of infection so that negative findings exclude the likelihood of meningitis in almost all cases.63,65,66 Patients with acute bacterial meningitis often demonstrate marked abnormalities in the chemistry analysis of the CSF, with protein concentrations of >100 mg/dL and glucose concentrations <45 mg/dL (or a CSF/blood glucose ratio of <0.5) due, in part, to disruption of the blood–brain barrier.63,66

Hematologic analysis of the CSF involves measurement of the WBC count with corresponding differential. Patients with acute bacterial meningitis often demonstrate an elevated CSF WBC count (>1,000 cells/mm3) with a neutrophilic predominance (>80% neutrophils). In contrast, patients with viral, fungal, or mycobacterial meningitis often display lower CSF WBC counts (5 to 1,000 cells/mm3) with a predominance of lymphocytes. In cases of a traumatic lumbar puncture (surrounding blood vessels are damaged during needle insertion), peripheral blood can enter the subarachnoid space and contaminate the CSF, making interpretation of the CSF WBC difficult. When interpreting the CSF WBC count after a traumatic tap, there should be one WBC for every 500 to 1,000 RBCs (based on blood composition); this ratio should be used to calculate a corrected CSF WBC during CSF analysis/interpretation.

Cerebrospinal Fluid Stain and Culture

For patients with suspected bacterial meningitis, a Gram stain and culture should be performed on CSF. Gram stain will demonstrate an organism in 60% to 90% of patients with bacterial meningitis and is helpful in selecting appropriate empiric antibiotic therapy.2,63,65-67 However, the sensitivity of the CSF Gram stain diminishes to 40% to 60% in patients who have received antibiotics prior to the lumber puncture (also known as partially treated meningitis).2,65 In patients with meningitis caused by viruses, fungi, or mycobacteria, the Gram stain is usually negative, and specialized tests should be used, such as the India ink stain or cryptococcal antigen test for the detection of Cryptococcus neoformans or the acid-fast stain for the detection of Mycobacterium tuberculosis.

All CSF specimens should be processed for culture based on the type of meningitis (acute versus chronic) and the organism suspected of causing the infection. In patients with bacterial meningitis, the cultures are often positive within 24 to 48 hours. In patients with nonbacterial meningitis, culture specimens should be incubated for longer periods of time (up to 2 to 6 weeks), because some organisms take longer to grow.

Other Specialized Tests

Several specialized tests may be performed on CSF specimens to aid in the detection of the causative organism, including bacterial antigen detection using LA, latex fixation, or enzyme immunoassay (EIA); fungal antigen detection; antibody detection; and bacterial, viral, or mycobacterial PCR assays.63-67

Bacterial antigen testing on CSF specimens is a rapid diagnostic test with results available within 10 to 15 minutes. Commercially available tests use antibody-coated particles that bind to specific capsular antigens of the most common pathogens that cause acute bacterial meningitis, including S pneumoniae, N meningitidis, H influenzae type B, and group B streptococci. The tests are performed by combining CSF (although they can also be performed using urine or serum) with antibody-coated particles and observing for agglutination, which signifies the presence of the bacterial antigen in the specimen. If visible agglutination does not occur, either the antigen is not present or it is present in insufficient amounts to cause detectable agglutination. Routine bacterial antigen detection of CSF specimens is not currently recommended because the results lack high specificity/sensitivity (not better than the traditional Gram stain), have an inadequate negative predictive value, and their use has rarely impacted patient treatment or been demonstrated to be cost-effective.3,63,67 However, bacterial antigen testing may be useful in patients with negative Gram stains or in patients who have received previous antimicrobial therapy.2,63-65,67

Nucleic-acid amplification tests, including PCR, are rapid and accurate tests for the diagnosis of meningitis due to bacteria, viruses, and fungi.66,67 CSF PCR results are often positive early in the course of infection and even remain positive during the first week of therapy.66 Several commercial PCR assays are available that amplify small amounts of specific DNA of the target organism followed by subsequent identification and verification. The FilmArray meningitis/encephalitis panel (BioFire Diagnostics, Salt Lake City, UT) is a multiplex PCR with high sensitivity of 94.2% and specificity of 99.8% that rapidly (<1 hr) detects 14 common causes (six bacteria [Escherichia coli K1, Haemophilus influenzae, Listeria monocytogenes, Neisseria meningitidis, Streptococcus agalactiae, Streptococcus pneumoniae]; seven viruses [Cytomegalovirus, Enterovirus, Herpes simplex virus 1, Herpes simplex virus 2, Human herpesvirus 6, Human parechovirus, Varicella zoster virus], and Cryptococcus neoformans) of meningitis/encephalitis directly from CSF.66,67

Streptococcal Pharyngitis

Acute pharyngitis is one of the most common infections encountered in medicine and can occur in both children and adults. Acute pharyngitis can be caused by several organisms (eg, bacteria and viruses), which produce similar signs and symptoms of infection. Antibiotic therapy is recommended only for patients with pharyngitis caused by bacteria, especially group A streptococci (Streptococcus pyogenes).68 Because group A strep pharyngitis comprises only a small percentage (20% to 30%) of patients with acute pharyngitis, it is important that a rapid, reliable diagnostic test be available to avoid unnecessary antibiotic use in patients with acute viral pharyngitis.68

The gold standard diagnostic test for acute pharyngitis caused by group A streptococcus is the throat culture, which often takes 1 to 2 days for results. Therefore, rapid antigen detection tests (RADTs) have been developed to expedite and confirm the diagnosis of group A streptococcal pharyngitis, with most tests yielding results within 15 minutes.68,69 Positive RADT test results expedite the initiation of antibiotic treatment in the appropriate patient. Several RADT tests are commercially available, with the newer tests employing EIA or chemiluminescent DNA probes (>95% specificity and ≥90% sensitivity).68,69 There are limited studies comparing the performance of different RADT tests to throat culture (the gold standard), so current recommendations suggest that traditional throat culture be performed in children and adolescents with a negative RADT test result to definitively exclude group A streptococcal pharyngitis.68,69

Pneumonia

Several obstacles make the diagnosis of bacterial pneumonia quite difficult. First, the respiratory tract is colonized with bacteria that may or may not be contributing to the infectious process. When obtaining a sample for culture, lower respiratory tract secretions can become contaminated with secretions or bacteria colonizing the upper respiratory tract; therefore, expectorated sputum samples should be evaluated to determine if contamination with saliva or upper respiratory tract flora has occurred (assessing of the adequacy of the sample).1,2,4,6,7,70,71 If bacteria other than normal respiratory flora are isolated, the clinician must determine the relative importance and significance of the organism(s) as a potential cause of pneumonia, in addition to assessing the patient for signs and symptoms of pneumonia. It is estimated that 40% to 60% of hospitalized patients with CAP are unable to produce a sputum sample; and 40% to 60% of produced samples that are submitted are judged as being inadequate.70 For this reason, in some patients, adequate sputum specimens are difficult to obtain without invasive procedures. Invasive procedures, such as BAL or protected specimen brush (PSB), are occasionally used to aid in the diagnosis of pneumonia in patients who are unable to expectorate an adequate sputum sample (especially in patients not responding to appropriate empiric therapy), in immunocompromised patients, and in patients with hospital-acquired pneumonia (HAP) or ventilator-associated pneumonia (VAP).2,19,70,71 Despite the best efforts at obtaining a lower respiratory tract sputum specimen for culture, as many as 30% to 50% of patients with pneumonia have negative culture results.70,71

To obtain an adequate expectorated sputum sample, the patient should be instructed to provide sputum generated from a deep cough. All expectorated sputum samples should be screened to ensure that the specimen is adequate and has not been contaminated by saliva or upper respiratory tract flora prior to processing for culture.2,70,71 Information used to assess the adequacy of an expectorated sputum sample is derived from visualization of the Gram stain of the specimen. Expectorated sputum specimens that contain >25 WBCs/hpf (unless the patient is neutropenic) and <10 squamous epithelial cells/hpf are considered adequate for further processing and culture.4,70,71 Samples with >10 epithelial cells/hpf are representative of upper respiratory tract contamination (saliva) and should not be processed for culture. The sputum Gram stain from an adequate sputum specimen may be used to guide empiric antibiotic therapy when the specimen is purulent and contains a predominant organism. Antibiotic therapy should be modified based on the culture results, especially if they reveal an infecting organism.

Because of the difficulty with collection and low yield with sputum culture, several rapid direct detection tests have been developed, including urinary antigen detection (Streptococcus pneumoniae and Legionella pneumophila serogroup 1) or NA-based methods on respiratory specimens.70,71 The S pneumoniae urinary antigen test may be useful in hospitalized patients who are unable to produce a sputum sample, in patients with severe pneumonia requiring intensive care unit admission, in patients at risk for pneumococcal pneumonia (eg, asplenic, alcohol abuse, liver disease), in patients with pneumonia and concomitant pleural effusion, and in patients who have received antibiotics before a specimen for culture has been obtained.70,71 Several NA-based rapid (in 1 hour) detection methods are commercially available for the detection of respiratory viruses and bacteria capable of causing upper and lower respiratory tract infections, and include tests such as the Verigene Respiratory Pathogens Flex Test (Luminex; detects three Bordetella spp. and 13 viral targets, including adenovirus, influenza, parainfluenza, rhinovirus, and RSV), the FilmArray Respiratory EZ Panel (BioFire Diagnostics; detects coronavirus, adenovirus, five influenza, human rhinovirus/enterovirus, parainfluenza, RSV, human metapneumovirus, B pertussis, M pneumoniae, and C pneumoniae), the FilmArray Respiratory Panel (BioFire Diagnostics; detects four coronaviruses, adenovirus, five influenza, human rhinovirus/enterovirus, four parainfluenza viruses, RSV, human metapneumovirus, B pertussis, B parapertussis M pneumoniae, and C pneumoniae), and the FilmArray Pneumonia Panel (BioFire Diagnostics; detects eight viruses, 18 bacteria associated with HAP and seven genetic markers of resistance).70 Additionally, serologic tests may also be used in the diagnosis of pneumonia caused by atypical bacteria such as L pneumophila, Mycoplasma pneumoniae, or Chlamydia pneumoniae because they are difficult to culture in the laboratory.2,3,70,71

In patients with HAP or VAP, semiquantitative analysis of tracheal aspirates or sputum cultures or quantitative analysis from BAL specimens may occasionally be performed to differentiate between infection and colonization based on the history of prior antibiotic use and the number of organisms recovered in the sputum specimen.19 Diagnostic thresholds for pneumonia based on colony counts recovered from a quantitative BAL specimen may differ among institutions. Studies evaluating quantitative BAL or PSB specimens for the diagnosis of HAP or VAP use a diagnostic threshold between 103 and 105 CFU/mL of an organism for the diagnosis of pneumonia.2,19,70

Genitourinary Tract Infections

Urinary Tract Infections

Urinary tract infections (UTIs) are common infections, prompting >8 million office visits and more than 100,000 hospitalizations per year.72-75 UTIs are especially common in female patients because of the close proximity of the urethra (which is shorter than in male patients) to the perirectal and vaginal regions, which are both colonized with bacteria. Because of this anatomic difference, bacteria are able to easily ascend the urethra in female patients and potentially cause infection in the bladder (cystitis) and upper urinary tract (pyelonephritis). In addition, hospitalized patients (male and female) with indwelling urinary catheters are at increased risk for developing UTIs, with approximately 20% of catheterized patients developing a UTI, even with only short-term catheterization.73,76

Under normal circumstances, urine within the bladder is sterile because all anatomic sites within the urinary tract above the urethra are not colonized with bacteria. However, the urethra is colonized with bacteria. If noninvasive urine collection methods are used for specimen collection, urine travels through the urethra and may inadvertently collect bacteria while passing through this nonsterile environment. Therefore, diagnostic criteria have been developed to discriminate between infection, bacterial colonization, or bacterial contamination based on quantitative bacterial colony counts from urine cultures and the presence of inflammatory cells and epithelial cells in the urinalysis.72-74

Urine samples for urinalysis and culture can be collected several ways. The most common method involves the collection of a clean-catch, midstream urine sample. Before obtaining the sample, the patient should be instructed to clean and rinse the periurethral area with a mild detergent and then retract the labial folds or penile foreskin when beginning to urinate. The patient should attempt to collect the urine in a sterile cup at the midpoint of the urine stream, collecting the urine sample a few seconds after the start of urination.

Other methods for specimen collection involve invasive procedures, such as obtaining urine via bladder catheterization (straight catheter) or via suprapubic bladder aspiration. Both of these methods avoid the potential contamination of the urine specimen by the urethra because the urine is collected directly from the bladder. In hospitalized patients with short-term indwelling urinary catheters, urine specimens should be collected directly from the urinary catheter by aspirating the catheter port or tubing (representing freshly voided urine) rather than obtaining the specimen from the collection bag (urine collected over a period of time).4,72,73,76 In patients with long-term indwelling urinary catheters, the catheter should be exchanged with the urine sample collected upon insertion of the new catheter.76 In all cases, urine samples should be immediately transported to the laboratory for processing.

Urine samples from women with acute uncomplicated cystitis are usually only evaluated using screening tests because the results are rapidly available and are useful at excluding the presence of a UTI.72 The most common rapid screening tests include commercially available reagent test strips, or urine dipsticks, that contain the leukocyte esterase test and the nitrate reductase test, and provide a negative predictive value of 98%.72,73 The leukocyte esterase test detects the presence of leukocyte esterase, which is an enzyme found in neutrophils. The nitrate reductase test detects the presence of urinary nitrite produced by the reduction of nitrate by nitrate-reducing enzymes of common urinary tract pathogens (primarily Enterobacterales).72-74 Positive results from either the leukocyte esterase test or nitrate reducatase test lead to initiation of treatment for a UTI without the need for urine culture in women with symptoms suggesting acute uncomplicated cystitis.

The urine from patients with recurrent UTIs, complicated/upper tract UTIs, or catheter-associated UTIs is typically evaluated using a urinalysis (microscopic examination) and urine culture. The urinalysis is a rapid test that involves the macroscopic and microscopic examination of the urine sample for color, clarity, specific gravity, and the presence of protein, glucose, RBCs, WBCs, bacteria, and epithelial cells. The urinalysis is performed either manually or with automated instruments. Urinalysis findings suggestive of a UTI include specimen cloudiness and the presence of pyuria (>10 WBC/mm3).72-74 The detection of pyuria, hematuria, proteinuria, or bacteriuria in the urinalysis may be an indication of infection, but none of these alone is specific for infection. The presence of squamous epithelial cells (>2 to 5 epithelial cells/mm3) in a urine sample suggests poor specimen collection and possible contamination.

The urine culture remains the hallmark laboratory test for the diagnosis of UTIs, with quantitative cultures providing the most useful data for determining the clinical significance of isolated bacteria. To establish the diagnosis of a UTI, urine cultures from midstream urine samples should display >105 CFU/mL of a single potential uropathogen with concomitant pyuria on urinalysis; however, some women with symptomatic cystitis may have lower colony counts of bacteria (103).73 Colony counts of >103 CFU/mL with pyuria are considered clinically relevant in urine specimens from patients with indwelling urethral catheters or intermittent catheterization, from men, or from children.72,75 Urine specimens obtained by suprapubic aspiration that display >102 CFU/mL with pyuria are indicative of the presence of infection.72,73,76

Prostatitis

Bacterial prostatitis can present as an acute or chronic infection that typically occurs in men >30 years of age.74 The diagnosis of acute bacterial prostatitis is often based on clinical presentation and the presence of bacteria in a urine specimen. Digital palpation of the prostate and prostatic massage to express purulent secretions are not recommended for the diagnosis of acute bacterial prostatitis because it may induce bacteremia. Conversely, the diagnosis of chronic bacterial prostatitis often cannot be established based on clinical grounds alone because the symptoms are nonspecific and the prostate is often not acutely inflamed. Therefore, the diagnosis of chronic prostatitis is classically established through the analysis of sequential urine and prostatic fluid cultures.74,77 Initially, two samples of urine are obtained for culture—one sample on initiation of urination (VB-1) and one sample obtained at midstream (VB-2). Next, prostate fluid is obtained for culture by massaging the prostate to produce expressed prostatic secretions. Lastly, a urine sample (VB-3) is obtained after prostatic secretions have been obtained and sent for culture. The diagnosis of chronic bacterial prostatitis is made when the expressed prostatic secretion sample contains > 10 times the quantity of bacteria cultured from VB-1 or VB-2 or if the VB-3 contains 10 times the quantity of bacteria cultured from VB-1 or VB-2.74,77

Sexually Transmitted Diseases

Gonorrhea

Infection due to N gonorrhoeae is the second most common notifiable sexually transmitted disease (STD) reported in the United States, with most infections involving the mucosa of the cervix, the urethra, the rectum, and the pharynx.78,79 Infections caused by N gonorrhoeae include localized, uncomplicated, or complicated genital infections (eg, urethritis, cervicitis, endometritis, pelvic inflammatory disease [PID] in women, and urethritis or epididymitis in men), pharyngitis, anorectal infections, and disseminated infection (eg, septic arthritis and meningitis) in both men and women.78,79 Women with genital tract infection and patients with pharyngeal infection are often asymptomatic, while men with urethritis often display symptoms of dysuria and urethral discharge. In addition, patients with N gonorrhoeae are often coinfected with other STDs, such as Chlamydia trachomatis, syphilis, or Trichomonas vaginalis; therefore, the diagnosis and treatment of all possible STDs in the patient and their sexual partners are important public health considerations in the control of STDs.78

The diagnosis of infection due to N gonorrhoeae can be established using Gram-stained smears, culture, or nonculture techniques detecting cellular components (only NA amplification tests [NAATs] are currently recommended for routine use; EIA and DNA probe tests are no longer recommended) of urethral, endocervical, or urine specimens (NAAT only) that detect cellular components of N gonorrhoeae.4,78-81

A presumptive diagnosis of gonorrhea can be made using direct microscopic examination of a clinical specimen using a Gram stain and oxidase test, in which gram-negative, oxidase-positive diplococci are demonstrated.78-81 In addition, the presence of neutrophils on a Gram stain of a urethral specimen is also helpful in establishing the presumptive diagnosis of urethritis.79 The Gram stain is both sensitive and specific for the presumptive diagnosis of N gonorrhoeae as a point-of-care test for symptomatic men with urethral discharge but is not as useful as a single diagnostic test in asymptomatic men or when evaluating endocervical or pharyngeal specimens.78,79 Additional tests, such as culture, should be performed to confirm the identification of the organism.

Culture on selective media remains the diagnostic standard for the identification of N gonorrhoeae.79,80 Culture is recommended for the diagnosis of gonorrhea from urethral, endocervical, vaginal, pharyngeal, or rectal swab (plastic or wire shafts with rayon, Dacron, or calcium alginate tips) specimens and should be performed on specimens from all patients (and sexual partners) with suspected gonococcal infections.79 Culture is also used as a confirmatory test in patients who have suspected gonorrhea based on positive Gram-stained smears or nonculture tests if the specimen has been adequately maintained. However, culture is not optimal in all circumstances due to the tenuous viability of the organism during storage and transport, which is what led to the development of nonculture tests for the detection of gonorrhea.79 Occasionally, susceptibility testing is performed on N gonorrhoeae isolates, especially in patients with suspected or documented treatment failure, to guide the choice of antibiotic therapy, as well as for epidemiologic purposes.78 In either case, patients are typically given empiric therapy with an antibiotic that demonstrates excellent activity against gonorrhea, keeping in mind that the incidence of β-lactamase-producing, penicillin-resistant gonococci is increasing.

In many clinical settings, nonculture tests for the detection of N gonorrhoeae have replaced traditional culture. Nonculture tests include NAATs, which are able to amplify organism-specific DNA sequences, and the NA hybridization (probe) test, which hybridizes any complementary rRNA that is present in the specimen (cannot differentiate organisms).78,79 Several NAATs for the detection of N gonorrhoeae are commercially available and have been designed to detect RNA or DNA sequences using amplification techniques (even on nonviable organisms). These tests have been FDA-approved for the detection of N gonorrhoeae in endocervical and vaginal swabs from women, urethral swabs from men, and urine samples from men and women; because the tests are different, the product information for each individual test should be consulted to dictate the collection methods and clinical specimen type that is suitable for each test.78,79 These tests are also useful for the detection of N gonorrhoeae from clinical specimens that have not been adequately maintained during transport or for collection for culture methods to be used. The major drawback of nonculture techniques for the detection of gonorrhea is that they cannot provide information on antibiotic susceptibility of the organism.81

Chlamydia

Chlamydia trachomatis is the most frequently reported infectious disease in the United States, with >1.3 million cases reported to the Centers for Disease Control and Prevention in 2010.78,79 Infection with C trachomatis is now a reportable communicable disease in the United States, with the highest prevalence in persons aged ≤24 years.78 C trachomatis can cause a number of infections, including cervicitis, endometritis, and PID in women; and urethritis, epididymo-orchitis, prostatitis, and proctitis (via receptive anal intercourse) in men.78 Infection with C trachomatis is also thought to contribute to female infertility and ectopic pregnancies. It is estimated that more than $500 million is spent annually on the direct costs associated with the management of C trachomatis infections.79

Most patients with chlamydial infections are asymptomatic, so screening is necessary to detect the presence of the organism.78,79 Because of the asymptomatic nature of chlamydia, it is thought that the current rates of reporting underestimate the true incidence of infection due to this organism. Chlamydia screening is now recommended annually in all sexually active women aged <25 years and other women at increased risk for infection (eg, new sexual partner, multiple sexual partners, sexual partner with an STD).78 In addition, chlamydia screening is also recommended in patients with other STDs because chlamydia often coexists with other STD pathogens.

Culture and nonculture methods are available for the detection of chlamydia. Culture involves the inoculation of the biologic specimen onto a confluent monolayer of cells that support the growth of C trachomatis. The culture is evaluated at 24 to 72 hours for the presence of intracellular inclusions using a fluorescent monoclonal antibody stain, which occur as a result of C trachomatis infection.79 Cell culture is not routinely used by most laboratories because of lack of standardization, technical difficulty, cost, and length of time to yield results (at least 48 hours). Therefore, other nonculture approaches for the laboratory diagnosis of chlamydia have been developed, including the direct fluorescent antibody (DFA) test and NAATs.80

Direct fluorescent antibody testing involves the staining of a biologic specimen with a fluorescein-labeled monoclonal antibody that binds to C trachomatis-specific antigens (elementary bodies).80 If the patient is infected with C trachomatis, the antibodies will react with the elementary bodies of the chlamydia in the secretions to produce fluorescence. DFA tests require significant technologist time for performance, so they are typically only used as a confirmatory test to other antigen detection tests.80

The other nonculture test used for the detection of C trachomatis is the NAAT, which has largely replaced tissue culture and DFA testing because of greater sensitivity and specificity.78,79,81 Several commercially available NAATs for the detection of C trachomatis have been designed to detect RNA or DNA sequences using PCR, ligase chain reaction, and various amplification techniques. These tests have been FDA approved for the detection of C trachomatis in endocervical or vaginal swabs from women, urethral swabs from men, and rectal swab or first catch urine samples from men and women.78,79

Syphilis

The spirochete T pallidum is the causative pathogen of an STD known as syphilis. There are a number of clinical manifestations and stages of syphilis that are based primarily on presenting symptoms and the natural history of the infection46,78,81:

  1. Primary syphilis: characterized by painless ulcers called chancres that are typically located at the site of inoculation or initial infection (usually in genital area) and spontaneously resolve over 1 to 8 weeks.

  2. Secondary syphilis: characterized by systemic symptoms, including fever, weight loss, malaise, headache, lymphadenopathy, and a mucocutaneous skin rash (generalized or localized, often involving the palms or the soles of the feet) resulting from hematogenous or lymphatic spread of the organism. If untreated, the manifestations resolve within 4 to 10 weeks.

  3. Latent syphilis: occurs after secondary syphilis, in which the organism is still present, but the patient is without symptoms. Latent syphilis acquired within the preceding year is categorized as early latent syphilis, whereas syphilis acquired more than 1 year ago or of unknown duration is categorized as late latent syphilis. This subclinical infection can be detected only by serologic tests.

  4. Late/tertiary syphilis and neurosyphilis: occurs in approximately 35% of untreated patients up to 10 to 25 years after initial infection; clinical manifestations are caused by progressive inflammatory disease that can involve the CNS (categorized as neurosyphilis) or outside the CNS (referred to as tertiary syphilis) including cardiovascular lesions (ascending aorta) and granuloma-like lesions (gummas) in the skin, bone, or visceral organs.

T pallidum cannot be grown in culture; therefore, the diagnosis of syphilis involves the direct detection of the spirochete in biologic specimens by microscopy or the detection of treponemal-specific antibodies using serologic testing.

Direct detection methods can be performed on appropriate clinical specimens obtained from suspicious genital or skin lesions, including lesion exudate or tissue. The direct detection of T pallidum using dark-field microscopy involves the immediate examination (within 20 minutes of collection) of the biologic specimen under a microscope with a dark-field condenser, looking for the presence of motile spirochetes, where T pallidum appears as 8- to 10-μm spiral-shaped organisms.4,46,81 Another test for the direct detection of T pallidum is the direct fluorescent antibody (DFA-TP) test in which the biologic specimen is combined with fluorescein-labeled monoclonal or polyclonal antibodies specific for T pallidum and examined by fluorescence microscopy.46,78 The interaction between the antibodies and treponemal-specific antigens produces fluorescence that can be visualized using microscopy.

There are two types of serologic tests that are used for the diagnosis of syphilis – nontreponemal and treponemal antibody tests. The use of only one type of serologic test is not sufficient for the diagnosis of syphilis (may result in false-negative or false-positive diagnoses), so persons with a reactive nontreponemal test result should undergo treponemal antibody testing to confirm the diagnosis.78

The nontreponemal antibody tests include the Venereal Disease Research Laboratory (VDRL) test and the rapid plasma reagin (RPR) test.46,81 Both the VDRL and RPR measure the presence of reagin, an antibody-like protein produced in patients with syphilis. However, reaginic antibodies are also produced in patients with other infections and conditions including autoimmune diseases, leprosy, TB, malaria, pregnancy, and injection drug use, so false-positive RPR results may occur.46,81 Both the RPR and VDRL tests are flocculation tests in which visible clumps are produced in the presence of the reagin antibody (T pallidum) in the submitted specimen. For the VDRL test, the biologic specimen (serum, CSF) is combined with cardiolipin-lecithin coated cholesterol particles on a glass slide and examined microscopically.46 If the reagin antibody is present in the biologic specimen, visual clumping occurs and is reported as reactive (medium and large clumps). The VDRL can be performed on serum and CSF as a quantitative test in which dilutions of the biologic specimen can be evaluated for reactivity; the dilution that produces a fully reactive result is reported as the VDRL titer (eg, 1:8 or 1:32). Therefore, the VDRL titer can be used to monitor a patient’s response to therapy. The high titers present in untreated disease (eg, 1:32) traditionally decrease 4-fold within 6 to 12 months of treatment and become undetectable in 1 to 2 years.

The RPR test is a modification of the VDRL test and is commercially available as a reaction card. Serum from the patient is placed on the reaction card and observed for clumping. The RPR result is quantified by evaluating dilutions of the biologic specimen for reactivity, with the highest dilution that produces a fully reactive result being reported as the RPR titer (eg, 1:8 or 1:32). The RPR titer is also used to monitor a patient’s response to therapy, where a 4-fold decline in titer 6 to 12 months after therapy would be suggestive of response. The RPR is easier to perform than the VDRL and is used by many laboratories and blood banks for routine syphilis screening. However, the RPR should not be used for the analysis of CSF specimens.

The nontreponemal antibody detection tests are nonspecific, so they are most useful for screening for the presence of syphilis.46,81 Because it takes several weeks for the development of reagin antibodies after exposure to syphilis, false-negative results (up to 25% of patients with primary syphilis) can occur in the early stages of the disease. In addition, false-positive results (up to 1% to 2%) can also occur because of the numerous other conditions where reagin antibodies are produced.3,46 Therefore, a positive RPR or VDRL test result should be confirmed with the fluorescent treponemal antibody absorption (FTA-ABS) or the microhemagglutination T pallidum (MHA-TP) test, both of which measure the presence of treponemal-specific antibodies.

The other type of serologic test measures the presence of treponemal antibodies and includes the FTA-ABS test and the MHA-TP test.46,78,81 In the FTA-ABS test, the patient’s serum or CSF is initially absorbed with non-T pallidum antigens to reduce cross-reactivity and then applied to a slide on which T pallidum organisms have been fixed followed by addition of a fluorescein-conjugated antihuman antibody for detection of specific antitreponemal antibodies. The amount of fluorescence is subjectively measured by the laboratory technician and reported as reactive, minimally reactive, or nonreactive. Therefore, this test is difficult to standardize among different laboratories. Because this test is also fairly expensive, it is primarily used to verify the results of a positive VDRL or RPR, rather than as a routine screening tool.46,81 The FTA-ABS test can detect antibodies earlier in the course of syphilis than nontreponemal tests and, once positive, remains positive for the life of the patient.

The MHA-TP test is performed using erythrocytes from a turkey, sheep, or other mammal that have been coated with treponemal antigens. These erythrocytes are then mixed with the patient’s serum and observed for agglutination, which signifies the presence of antibodies directed against T pallidum. The results are reported as reactive (positive) or nonreactive (negative). Lastly, EIA tests and PCR-based tests for the detection of T pallidum are being evaluated as screening or confirmatory tests for the diagnosis of syphilis, especially for patients in whom serologic testing is not reliable.46,81

Trichomonas

Infection caused by the protozoan T vaginalis is the most common, nonviral STD in the United States, affecting 3.7 million people.78 T. vaginalis is typically diagnosed through detection of actively motile organisms during microscopic examination of wet mount preparations of vaginal secretions, urethral discharge, prostatic fluid, or urine sediment.61,78,81 Because the sensitivity of the wet mount preparation is 50% to 80%, other diagnostic tests have been developed for the detection of T vaginalis to enhance diagnostic yield, sensitivity, and specificity.78,81 Culture using Diamond’s medium is considered the diagnostic gold standard test because it is associated with >80% sensitivity; however, culture methods require proper collection and rapid inoculation for best results, so it is not routinely performed by most laboratories.81 Several rapid antigen detection methods are commercially available for the diagnosis of infection caused by T vaginalis that are easy to perform and employ different assays (IFA and capillary flow ICA).61 Lastly, NA detection methods are highly sensitive and specific tests for the detection of Trichomonas and include direct DNA probe Affirm VPIII (BD) and the APTIMA T vaginalis Assay (Hologic-GenProbe, San Diego, CA).61,78

Assessing Sterile Body Fluids for Presence of Infection

Sterile body fluids such as pericardial fluid (pericarditis), pleural fluid (empyema), synovial fluid (septic arthritis), and peritoneal fluid (peritonitis) can be analyzed for the presence of infection. The specimens should be aseptically obtained by needle aspiration, placed in sterile collection tubes, and immediately transported to the laboratory for fluid analysis and culture. Approximately 1 to 5 mL of fluid should be obtained when analyzing pericardial, pleural, or synovial fluid, while up to 10 mL of peritoneal fluid is required for the diagnosis of peritonitis.82 All sterile fluids should be processed for cell count (establishing the presence of WBCs with differential), chemistry (protein and glucose), direct microscopic examination including Gram stain (presence of bacteria), and culture. For pleural and synovial fluids, specific criteria are available to aid in the diagnosis of infection (Tables 18-13 and 18-14).82-88 Peritoneal fluid characteristics that may be suggestive of peritonitis include a WBC of >250 cells/mm3, a lactate concentration >25 mg/dL, a pH <7.35, a fluid/blood glucose ratio of <0.7 (in TB peritonitis), and an elevated protein concentration (except in patients with cirrhosis).89,90 The diagnosis of infection in each of these sites should be established based on the presence of WBCs and other characteristic chemistry abnormalities in the sterile fluid specimen, the growth of a pathogenic organism from the cultured material, and the characteristic signs and symptoms of infection at that site.

TABLE 18-13.

Pleural Fluid Findings and Interpretation

TRANSUDATIVE (SUGGESTIVE OF CONGESTIVE HEART FAILURE, CIRRHOSIS)

EXUDATIVE (SUGGESTIVE OF INFECTION SUCH AS EMPYEMA, MALIGNANCY, PANCREATITIS WITH ESOPHAGEAL PERFORATION, SLE)

Appearance

Clear, serous

Cloudy

pH

>7.2

<7.2

LDH (International Units/L)

<200

≥200

Pleural fluid LDH to serum LDH ratio

<0.6

>0.6

Protein (g/dL)

<3

>3

Pleural fluid to serum protein ratio

<0.5

>0.5

Glucose (mg/dL)

>60 (same as serum)

<40–60

WBCs (count/mm3)

<10,000

>10,000

WBC differential

<50% PMNs

If infectious, depends on pathogen

LDH = lactate dehydrogenase; PMNs = polymorphonuclear leukocytes; SLE = systemic lupus erythematosus.

Source: References 8285.
TABLE 18-14.

Synovial Fluid Findings and Interpretation

NORMAL

NONINFLAMMATORY (OSTEOARTHRITIS, TRAUMA, AVASCULAR NECROSIS, SLE, EARLY RHEUMATOID ARTHRITIS)

INFLAMMATORY (RHEUMATOID ARTHRITIS, SPONDYLOARTHROPATHIES, VIRAL ARTHRITIS, CRYSTAL-INDUCED ARTHRITIS)

PURULENT (BACTERIAL INFECTION, TUBERCULOUS INFECTION, FUNGAL INFECTION)

WBCs (count/mm3)

<150–200

<2,000

2,000–50,000

>50,000

WBC differential

No predominance

<25% PMNs

>70% PMNs, variable

>75% to 90% PMNs

Protein (g/dL)

1.3–1.8

3–3.5

>3.5

>3.5

Glucose (mg/dL)

Normal

Normal

70–90

<40–50

PMNs = polymorphonuclear leukocytes; SLE = systemic lupus erythematosus.

Source: References 8688.

ACUTE PHASE REACTANTS AND INFECTION

Chapters 16 and 20 provide information on the background, normal range, and clinical use of acute phase reactants such as the ESR, CRP, and procalcitonin in the diagnosis of inflammatory diseases. The ESR and CRP may also be elevated in the presence of infection.91-96 Elevations in the ESR and CRP do not differentiate between inflammatory or infectious processes because they increase in response to tissue injury of any cause. However, the ESR and CRP are often elevated in the presence of infection, with increased levels reported in bacterial otitis media, osteomyelitis, endocarditis, PID, septic arthritis, prosthetic joint infections, and infections in transplant patients, and they may serve as an adjunctive modality to aid in the diagnosis of these infections.91-96 ESR levels of >100 mm/hr have a high specificity for the presence of infection, malignancy, or arteritis.92 Serial measurement of the ESR, and especially the CRP, may also be useful in assessing the response to antibiotic therapy in the treatment of deep-seated infections such as endocarditis or osteomyelitis.91-96

Procalcitonin is the precursor of calcitonin, a calcium regulatory hormone, which is also an acute phase reactant that is produced in response to systemic inflammation.91,97-99 During infection, the metabolism of procalcitonin is altered in response to toxins and cytokines from bacteria, malaria, and some fungi (not viruses).97-100 As procalcitonin accumulates, its levels become detectable within 2 to 4 hours of infection and peak at 6 to 24 hours, with the extent of production correlating with bacterial load and severity of infection.97,98 It was originally believed that procalcitonin levels increased in response to tissue injury or sepsis induced only by infection; however, levels of procalcitonin may be elevated in other inflammatory diseases or situations, such as autoimmune diseases, severe trauma, cirrhosis, pancreatitis, burns, cardiac surgery, cardiac arrest, certain types of cancer, receipt of some conditioning agents prior to stem cell transplantation, and hypotension during surgery.91,92,97-100

The use of procalcitonin in the diagnosis of infection has been evaluated in numerous studies involving different patient types, infections, and clinical settings, with several procalcitonin-based algorithms being developed to (1) determine the presence of infection/guide initiation of antibiotic therapy, (2) evaluate the efficacy of empiric antibiotic therapy, and (3) determine when antibiotic therapy can be deescalated or discontinued during the treatment of an infection.97,98 Procalcitonin levels that correlate with the presence of infection have not been clearly defined for all infection types and clinical settings, but it does appear as if procalcitonin levels <0.1 mcg/L (eg, undetectable) exclude the presence of infection.98 In the past decade, the role of procalcitonin has been studied in the context of sepsis and critically ill patients, lower respiratory tract infections, chronic obstructive pulmonary disease, acute infections in the CNS, infectious complications of burns and pancreatitis, and polytrauma.101 These studies demonstrate a wide variability in diagnostic accuracy, usefulness in guiding antibiotic discontinuation, and cost effectiveness. Additional research is needed to further define the role of procalcitonin in the diagnosis and management of different patient types, infections, and clinical settings.97-99

SUMMARY

Although infectious disease is a rapidly changing field because of new challenges and technological advances, the diagnosis of infection is highly dependent on the proper performance and interpretation of numerous laboratory tests. For example, the Gram stain is a readily available, invaluable tool for examining clinical specimens for the presence of bacteria. Culture of clinical specimens using appropriate growth media allows for the cultivation and identification of many infecting bacteria, which often takes 24 to 48 hours. Numerous rapid diagnostic tests are now available for the identification of bacteria directly from clinical specimens, such as blood, stool, respiratory secretions, or body fluids, which substantially decrease the time to organism identification when compared with traditional bacterial culture and identification methods.3 Susceptibility tests for rapidly growing aerobic bacteria are commonly performed using an automated microdilution or a manual disk diffusion method. Bacterial susceptibilities to various antimicrobial agents are reported as S, I, and R. National standards for susceptibility testing are available and help guide the performance of the tests, the choice of antimicrobial agents to evaluate for susceptibility, and the reporting procedures of susceptibility tests by the clinical microbiology laboratory. Empiric antimicrobial therapy is typically chosen based on the suspected site and subsequent potential causative organisms of infection (using local or regional susceptibility information). Once the results of bacterial culture and susceptibility testing are available, antimicrobial therapy is de-escalated, if possible, to a more targeted (directed) regimen based on the susceptibility profile of the infecting organism in conjunction with patient-specific (eg, clinical condition, site of infection, drug allergies, and renal function) and infection-specific information.

Lastly, several infection types (eg, meningitis, UTIs) and certain pathogens (eg, B burgdorferi and L pneumophila) often require specialized laboratory testing to aid in the identification of the infecting organism. The clinician should be aware of the diagnostic tests currently available for these infections.

LEARNING POINTS

1. What methods for antimicrobial susceptibility testing are used by most microbiology laboratories in the United States, and how is this information conveyed to the clinician?

ANSWER: Microbiology laboratories often use several methods for antimicrobial susceptibility testing to accurately determine the activity of antibiotics against many different types of bacteria (eg, aerobic, anaerobic, and fastidious). However, most laboratories predominantly use automated broth microdilution methods (Vitek 2, MicroScan) that utilize commercially prepared, disposable microtiter trays or cassettes for antimicrobial susceptibility testing that can test the susceptibility of multiple antibiotics simultaneously while decreasing cost and labor. Although many microbiology laboratories perform rapid diagnostic tests for the identification of bacteria and the detection of pertinent resistant gene markers, antimicrobial susceptibility testing (using automated microdilution methods) is still being performed to determine the exact susceptibility and resistance profile of the infecting bacteria. The antimicrobial susceptibility results for each bacteria are compiled in a report that contains the following information: antibiotic tested, MIC or MIC range (especially with automated broth microdilution methods), and CLSI interpretive criteria (S, I, and R). These reports are usually located in the patient’s medical chart (electronic or paper) and in the hospital laboratory information system. Refer to Minicase 2 for a specific example of an antimicrobial susceptibility report.

2. Is the antibiotic with the lowest MIC on an individual susceptibility report always the best antibiotic choice in the treatment of an infection?

ANSWER: No, the antibiotic with the lowest MIC on an individual susceptibility report may not always be the best choice for the treatment of a specific infection. While the MIC value is an indication of the in vitro activity of the antibiotic against an organism, other issues must be considered when choosing an antibiotic once susceptibility results have returned (these are the same considerations used to establish MIC breakpoints). Each antibiotic has its own unique pharmacokinetic profile and recommended dosage range, resulting in unique serum and site concentrations. An antibiotic might display potent in vitro active against a particular organism but may be ineffective in vivo because of poor penetration to the site of infection. Thus, both the site of infection and achievable serum/site concentrations should be considered. Other things that should also be considered when selecting an antibiotic include the pharmacodynamic parameter that correlates with efficacy. The pharmacokinetic-pharmacodynamic relationship for each antibiotic-organism combination as well as the results of Monte Carlo simulations (analyses performed using population pharmacokinetic data of the antibiotic and bacteria MIC distribution data from susceptibility studies to evaluate the percentage of time the antibiotic will achieve adequate pharmacodynamic indices for the treatment of that organism in a simulated population), if available, should be reviewed to determine if optimal pharmacodynamic exposure will occur with the selected antibiotic based on the infecting organism’s MIC value. The results from clinical efficacy studies should also be considered when selecting a specific antibiotic—has the antibiotic demonstrated efficacy in the treatment of the infection in controlled clinical trials/published studies and, if so, what MICs were observed in patients with clinical success versus clinical failure? Lastly, other factors, such as patient characteristics (eg, pregnancy, comorbidities, allergies), drug characteristics (eg, administration schedule, route of therapy, cost) and hospital/insurance company formulary, should also be considered when selecting an antibiotic.

3. What are the major laboratory tests that are used in the diagnosis of UTIs, meningitis, pneumonia, and septic arthritis?

ANSWER: In patients with signs and symptoms suggestive of a complicated UTI, a urine sample (clean-catch midstream, catheterized specimen, suprapubic aspiration) is usually sent to the laboratory for microscopic analysis (urinalysis) and culture. In patients with signs and symptoms suggestive of meningitis, a lumbar puncture is performed to obtain CSF that is evaluated for general appearance, glucose concentration, protein concentration, WBC count, WBC differential, Gram stain, and culture. In addition, depending on the medical history of the patient, specialized tests may also be performed on the CSF. In patients with signs and symptoms suggestive of pneumonia, a sputum sample (expectorated, BAL, PSB) is submitted to the laboratory for adequacy evaluation, Gram stain and culture, and, occasionally, S. pneumoniae urinary antigen. In patients with suspected septic arthritis, a synovial fluid aspirate is analyzed for cell count (presence of WBCs with differential), chemistry (protein and glucose), direct microscopic examination including Gram stain (presence of bacteria), and culture. Patients with septic arthritis may also have an elevated ESR or CRP. Also, in all infections discussed in this chapter, patients may also exhibit leukocytosis and a left shift, which would be demonstrated on a CBC.

REFERENCES

  • 1.

    Specimen management. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Elsevier Inc; 2017:56-71.

  • 2.

    Miller JM, Binnicker MJ, Campbell S, et al.A guide to utilization of the microbiology laboratory for diagnosis of infectious diseases: 2018 update by the Infectious Diseases Society of America (IDSA) and the American Society for Microbiology (ASM). Clin Infect Dis. 2018;67(6):e1-e94.PubMed

    • Search Google Scholar
    • Export Citation
  • 3.

    Patel R. The clinician and the microbiology laboratory: test ordering, specimen collection, and result interpretation. In: Bennett JE, Dolin R, Blaser MJ, eds. Principles and Practice of Infectious Diseases. 9th ed. Philadelphia, PA: Elsevier, Inc.; 2020:194-210.

    • Search Google Scholar
    • Export Citation
  • 4.

    McElvania E, Singh K. Specimen collection, transport, and processing: Bacteriology. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:302-330.

    • Search Google Scholar
    • Export Citation
  • 5.

    Popescu A, Doyle RJ. The Gram stain after more than a century. Biotech Histochem. 1996;71(3):145-151.PubMed

  • 6.

    Role of microscopy. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Elsevier Inc; 2017:72-85.

  • 7.

    Woods GL, Walker DH. Detection of infection or infectious agents by use of cytologic and histologic stains. Clin Microbiol Rev. 1996;9(3):382-404.PubMed

    • Search Google Scholar
    • Export Citation
  • 8.

    Atlas R, Snyder J. Reagents, stains, and media: bacteriology. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:331-361.

    • Search Google Scholar
    • Export Citation
  • 9.

    Graman PS, Menegus MA. Microbiology laboratory tests. In: Betts RF, Chapman SW, Penn RL, eds. A Practical Approach to Infectious Diseases. 5th ed. Philadelphia, PA: Lippincott, Williams and Wilkins; 2003:929-956.

    • Search Google Scholar
    • Export Citation
  • 10.

    Bacteria. In: Wilborn JW, ed. Microbiology. Springhouse, PA: Springhouse Corporation; 1993:36-50.

  • 11.

    Werth BJ, Barber KE, Smith JR, Rybak MJ. Laboratory tests to direct antimicrobial pharmacotherapy. In: DiPiro JT, Yee GC, Posey L, et al., eds. Pharmacotherapy: A Pathophysiologic Approach. 11th ed. New York, NY: McGraw-Hill; 2020. https://accesspharmacy.mhmedical.com/content.aspx?bookid=2577&sectionid=219306547. Accessed June 29, 2020.

    • Search Google Scholar
    • Export Citation
  • 12.

    Traditional cultivation and identification. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis: Elsevier Inc; 2017:86-112.

    • Search Google Scholar
    • Export Citation
  • 13.

    Lagier JC, Edouard S, Pagnier I, et al.Current and past strategies for bacterial culture in clinical microbiology. Clin Microbiol Rev. 2015;28(1):208-236.PubMed

    • Search Google Scholar
    • Export Citation
  • 14.

    Nucleic acid-based analytical methods for microbial identification and characterization. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Elsevier Inc; 2017:113-143.

    • Search Google Scholar
    • Export Citation
  • 15.

    Pliakos EE, Andreatos N, Shehadeh F, et al.The cost-effectiveness of rapid diagnostic testing for the diagnosis of bloodstream infections with or without antimicrobial stewardship. Clin Microbiol Rev. 2018;31(3):e1-e22 10.1128/CMR.00095-17.PubMed

    • Search Google Scholar
    • Export Citation
  • 16.

    Nolte FS. Molecular microbiology. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:86-123.

    • Search Google Scholar
    • Export Citation
  • 17.

    Versalovic J, Highlander SK, Ganesh BP, Petrosino JF. The human microbiome. Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:254-267.

    • Search Google Scholar
    • Export Citation
  • 18.

    Reese RE, Betts RF. Principles of antibiotic use. In: Betts RF, Chapman SW, Penn RL, eds. A Practical Approach to Infectious Diseases. 5th ed. Philadelphia, PA: Lippincott, Williams and Wilkins; 2003:969-988.

    • Search Google Scholar
    • Export Citation
  • 19.

    Kalil AC, Metersky ML, Klompas M, et al.Management of adults with hospital-acquired and ventilator-associated pneumonia: 2016 Clinical Practice Guidelines by the Infectious Diseases Society of America and the American Thoracic Society. Clin Infect Dis. 2016;63:1-51.

    • Search Google Scholar
    • Export Citation
  • 20.

    Laboratory methods and strategies for antimicrobial susceptibility testing. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:177-204.

    • Search Google Scholar
    • Export Citation
  • 21.

    Clinical and Laboratory Standards Institute (CLSI). Development of In Vitro Susceptibility Testing Criteria and Quality Control Parameters; Approved Guideline. 5th ed. Wayne, PA: Clinical and Laboratory Standards Institute; 2018.

    • Search Google Scholar
    • Export Citation
  • 22.

    Peterson LR, Shanholtzer CJ. Tests for bactericidal effects of antimicrobial agents: technical performance and clinical relevance. Clin Microbiol Rev. 1992;5(4):420-432.PubMed

    • Search Google Scholar
    • Export Citation
  • 23.

    Simner PJ, Humphries R. Special phenotypic methods for detecting antibacterial resistance. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:1316-1347.

    • Search Google Scholar
    • Export Citation
  • 24.

    DeGirolami PC, Eliopoulos G. Antimicrobial susceptibility tests and their role in therapeutic drug monitoring. Clin Lab Med. 1987;7(3):499-512.PubMed

    • Search Google Scholar
    • Export Citation
  • 25.

    Jorgensen JH, Ferraro MJ. Antimicrobial susceptibility testing: a review of general principles and contemporary practices. Clin Infect Dis. 2009;49(11):1749-1755.PubMed

    • Search Google Scholar
    • Export Citation
  • 26.

    Clinical and Laboratory Standards Institute (CLSI). Performance Standards for Antimicrobial Susceptibility Testing. 30th ed. Wayne, PA: Clinical and Laboratory Standards Institute; 2020.

    • Search Google Scholar
    • Export Citation
  • 27.

    Clinical and Laboratory Standards Institute (CLSI). Methods for Dilution Antimicrobial Susceptibility Tests for Bacteria That Grow Aerobically; Approved Standard. 11th ed. Wayne, PA: Clinical and Laboratory Standards Institute; 2018.

    • Search Google Scholar
    • Export Citation
  • 28.

    Clinical and Laboratory Standards Institute (CLSI). Performance Standards for Antimicrobial Disk Susceptibility Tests; Approved Standard. 13th ed. Wayne, PA: Clinical and Laboratory Standards Institute; 2018.

    • Search Google Scholar
    • Export Citation
  • 29.

    Koeth LM, Miller LA. Antimicrobial susceptibility test methods: dilution and disk diffusion methods. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:1284-1299.

    • Search Google Scholar
    • Export Citation
  • 30.

    Pulidot MR, García-Quintanilla M, Martín-Peña R, et al.Progress on the development of rapid methods for antimicrobial susceptibility testing. J Antimicrob Chemother. 2013;68(12):2710-2717.PubMed

    • Search Google Scholar
    • Export Citation
  • 31.

    Clinical and Laboratory Standards Institute (CLSI)/National Committee for Clinical Laboratory Standards. Methods for Determining Bactericidal Activity of Antimicrobial Agents. Wayne, PA: Clinical and Laboratory Standards Institute; 1999.

    • Search Google Scholar
    • Export Citation
  • 32.

    Reller LB. The serum bactericidal test. Rev Infect Dis. 1986;8(5):803-808.PubMed

  • 33.

    Clinical and Laboratory Standards Institute (CLSI)/National Committee for Clinical Laboratory Standards. Methodology for the Serum Bactericidal Test. Wayne, PA: Clinical and Laboratory Standards Institute; 1999.

    • Search Google Scholar
    • Export Citation
  • 34.

    Bush K. Past and present perspectives on β-lactamases. Antimicrob Agents Chemother. 2018;62(10):1-18.PubMed

  • 35.

    Paterson DL, Bonomo RA. Extended-spectrum β-lactamases: a clinical update. Clin Microbiol Rev. 2005;18(4):657-686.PubMed

  • 36.

    Humphries RM, Bard JD. Susceptibility test methods: fastidious bacteria. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:1348-1376.

    • Search Google Scholar
    • Export Citation
  • 37.

    Thomson KS. Extended-spectrum β-lactamases, AmpC, and carbapenemase issues. J Clin Microbiol. 2010;48(4):1019-1025.PubMed

  • 38.

    Abbott AN, Fang FC. Molecular detection of antibacterial drug resistance. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:1420-1431.

    • Search Google Scholar
    • Export Citation
  • 39.

    van Belkum A, Rochas O. Laboratory-based and point-of-care testing for MSSA/MRSA detection in the age of whole genome sequencing. Front Microbiol. 2018;9:1-9.PubMed

    • Search Google Scholar
    • Export Citation
  • 40.

    Anaerobic bacteriology: overview and general laboratory considerations. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:499-512.

    • Search Google Scholar
    • Export Citation
  • 41.

    Schuetz AN, Carpenter DE. Susceptibility test methods: anaerobic bacteria. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:1377-1397.

    • Search Google Scholar
    • Export Citation
  • 42.

    Clinical and Laboratory Standards Institute (CLSI). Methods for Antimicrobial Susceptibility Testing of Anaerobic Bacteria; Approved Standard. 9th ed. Wayne, PA: Clinical and Laboratory Standards Institute; 2018.

    • Search Google Scholar
    • Export Citation
  • 43.

    Schuetz AN. Antimicrobial resistance and susceptibility testing of anaerobic bacteria. Clin Infect Dis. 2014;59(5):698-705.PubMed

  • 44.

    Bordetella pertussis, Bordetella parapertussis and related species. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:475-479.

    • Search Google Scholar
    • Export Citation
  • 45.

    Sanchez E, Vannier E, Wormser GP, Hu LT. Diagnosis, treatment and prevention of Lyme disease, human granulocytic anaplasmosis, and babeiosis: a review. JAMA. 2016;315(16):1767-1777.PubMed

    • Search Google Scholar
    • Export Citation
  • 46.

    The spirochetes. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:578-589.

  • 47.

    Brucella. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:470-474.

  • 48.

    Obligate intracellular and nonculturable bacterial agents. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:555-669.

    • Search Google Scholar
    • Export Citation
  • 49.

    McDonald LC, Gerding DN, Johnson S, et al.Clinical practice guidelines for Clostridium difficile infection in adults and children: 2017 update by the Infectious Diseases Society of America (IDSA) and the Society for Healthcare Epidemiology of America (SHEA). Clin Infect Dis. 2018;66(7):e1-e48.PubMed

    • Search Google Scholar
    • Export Citation
  • 50.

    Intestinal protozoa. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:629-669.

  • 51.

    Dumler JS, Madigan JE, Pusterla N, et al.Ehrlichioses in humans: epidemiology, clinical presentation, diagnosis, and treatment. Clin Infect Dis. 2007;45(suppl 1):S45-S51.PubMed

    • Search Google Scholar
    • Export Citation
  • 52.

    Biggs HM, Behravesh CB, Bradley KK, et al.Diagnosis and management of tickborne rickettsial diseases: Rocky Mountain spotted fever and other spotted fever group rickettsioses, ehrlichioses, and anaplasmosis, United States. MMWR Recomm Rep. 2016;65(2)(No. RR 2):1-44.PubMed

    • Search Google Scholar
    • Export Citation
  • 53.

    Makristathis A, Hirschl AM, Mégraud F, Bessède E. Review: diagnosis of Helicobacter pylori infection. Helicobacter. 2019;24(suppl 1):e12641.PubMed

    • Search Google Scholar
    • Export Citation
  • 54.

    Edelstein PH. Legionella. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:905-920.

    • Search Google Scholar
    • Export Citation
  • 55.

    Blood and tissue protozoa. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:670-690.

    • Search Google Scholar
    • Export Citation
  • 56.

    Waites KB, Bebear C. Mycoplasma and Ureaplasma. In: Carroll KC, Pfaller MA, Landry ML, et al., eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:1117-1136.

    • Search Google Scholar
    • Export Citation
  • 57.

    Opportunistic atypical fungus: Pneumocystic jirovecii. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:822-824.

    • Search Google Scholar
    • Export Citation
  • 58.

    Mejia R, Weatherhead J, Hotez PJ. Intestinal nematodes (roundworms). In: Bennett JE, Dolin R, Blaser MJ, eds. Principles and Practice of Infectious Diseases. 9th ed. Philadelphia, PA: Elsevier, Inc.; 2020:3436-3442.

    • Search Google Scholar
    • Export Citation
  • 59.

    White AC Jr, Coyle CM, Rajshekhar V, et al.Diagnosis and Treatment of Neurocysticercosis: 2017 Clinical Practice Guidelines by the Infectious Diseases Society of America (IDSA) and the American Society of Tropical Medicine and Hygiene (ASTMH). Clin Infect Dis. 2018;66(8):e49-e75.PubMed

    • Search Google Scholar
    • Export Citation
  • 60.

    Murat JB, Hidalgo HF, Brenier-Pinchart MP, et al.Human toxoplasmosis: which biological diagnostic tests are best suited to which clinical situations? Expert Rev Anti Infect Ther. 2013;11(9):943-956.PubMed

    • Search Google Scholar
    • Export Citation
  • 61.

    Protozoa from other body sites. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:691-702.

    • Search Google Scholar
    • Export Citation
  • 62.

    Clinical and Laboratory Standards Institute (CLSI). Analysis and Presentation of Cumulative Antimicrobial Susceptibility Test Data; Approved Guideline. 4th ed. Wayne, PA: Clinical and Laboratory Standards Institute; 2014.

    • Search Google Scholar
    • Export Citation
  • 63.

    Tunkel AR, Hartman BJ, Kaplan SL, et al.Practice guidelines for the management of bacterial meningitis. Clin Infect Dis. 2004;39(9):1267-1284.PubMed

    • Search Google Scholar
    • Export Citation
  • 64.

    Meningitis and other. infections of the central nervous system. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:965-975.

    • Search Google Scholar
    • Export Citation
  • 65.

    Koutsari C, Dilworth TJ, Holt J, et al.Central nervous system infections. In: DiPiro JT, Yee GC, Posey L, et al., eds. Pharmacotherapy: A Pathophysiologic Approach. 11th ed. New York, NY: McGraw-Hill; 2020. https://accesspharmacy.mhmedical.com/content.aspx?bookid=2577&sectionid=219306547. Accessed June 29, 2020.

    • Search Google Scholar
    • Export Citation
  • 66.

    Hasbun R, Tunkel AR. Approach to the patient with central nervous system infection. In: Bennett JE, Dolin R, Blaser MJ, eds. Principles and Practice of Infectious Diseases. 9th ed. Philadelphia, PA: Elsevier, Inc.; 2020:1176-1182.

    • Search Google Scholar
    • Export Citation
  • 67.

    Poplin V, Boulware DR, Bahr NC. Methods for rapid diagnosis of meningitis etiology in adults. Biomarkers Med. 2020;14(6):459-479.PubMed

  • 68.

    Shulman ST, Bisno AL, Clegg HW, et al.Clinical practice guideline for the diagnosis and management of group A streptococcal pharyngitis: 2012 update by the Infectious Diseases Society of America. Clin Infect Dis. 2012;55(10):1279-1282.PubMed

    • Search Google Scholar
    • Export Citation
  • 69.

    Leung AK, Newman R, Kumar A, Davies HD. Rapid antigen detection testing in diagnosing group A β-hemolytic streptococcal pharyngitis. Expert Rev Mol Diagn. 2006;6(5):761-766.PubMed

    • Search Google Scholar
    • Export Citation
  • 70.

    Infections of the lower respiratory tract. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:942-956.

    • Search Google Scholar
    • Export Citation
  • 71.

    Metlay JP, Waterer GW, Long AC, et al.Diagnosis and treatment of adults with community-acquired pneumonia. An official clinical practice guideline of the American Thoracic Society and Infectious Diseases Society of America. Am J Respir Crit Care Med. 2019;200(7):e45-e67.PubMed

    • Search Google Scholar
    • Export Citation
  • 72.

    Graham JC, Galloway A. The laboratory diagnosis of urinary tract infections. J Clin Pathol. 2001;54:911-919.PubMed

  • 73.

    Infections of the urinary tract. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:987-998.

    • Search Google Scholar
    • Export Citation
  • 74.

    Fernandez JM, Coyle EA. Urinary tract infections and prostatitis. In: DiPiro JT, Yee GC, Posey L, et al., eds. Pharmacotherapy: A Pathophysiologic Approach, 11th ed. New York, NY: McGraw-Hill; 2020. https://accesspharmacy.mhmedical.com/content.aspx?bookid=2577&sectionid=219306547. Accessed June 29, 2020.

    • Search Google Scholar
    • Export Citation
  • 75.

    Gupta K, Hooton TM, Naber KG, et al.International clinical practice guidelines for the treatment of acute uncomplicated cystitis and pyelonephritis in women: A 2010 update by the Infectious Diseases Society of America and the European Society for Microbiology and Infectious Diseases. Clin Infect Dis. 2011;52(5):103-120.PubMed

    • Search Google Scholar
    • Export Citation
  • 76.

    Hooton TM, Bradley SF, Cardenas DD, et al.Diagnosis, prevention, and treatment of catheter-associated urinary tract infection in adults: 2009 International Clinical Practice Guidelines from the Infectious Diseases Society of America. Clin Infect Dis. 2010;50(5):625-663.PubMed

    • Search Google Scholar
    • Export Citation
  • 77.

    Meares EM, Stamey TA. Bacteriologic localization patterns in bacterial prostatitis and urethritis. Invest Urol. 1968;5(5):492-518.PubMed

    • Search Google Scholar
    • Export Citation
  • 78.

    Centers for Disease Control and Prevention. Sexually transmitted diseases treatment guidelines, 2015. MMWR Recomm Rep. 2015;64(RR-03):1-137.PubMed

    • Search Google Scholar
    • Export Citation
  • 79.

    Centers for Disease Control and Prevention. Recommendations for the laboratory-based detection Chlamydia trachomatis and Neisseria gonorrhoeae—2014. MMWR Recomm Rep. 2014;63:1-19.

    • Search Google Scholar
    • Export Citation
  • 80.

    Genital tract infections. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis: Mosby Inc; 2017:999-1014.

  • 81.

    Duhon B, Burnett Y. Sexually transmitted diseases. In: DiPiro JT, Yee GC, Posey L, et al., eds. Pharmacotherapy: A Pathophysiologic Approach, 11th ed. New York, NY: McGraw-Hill; 2020. https://accesspharmacy.mhmedical.com/content.aspx?bookid=2577&sectionid=219306547. Accessed June 29, 2020

    • Search Google Scholar
    • Export Citation
  • 82.

    Normally sterile body fluids, bone and bone marrow, and solid tissues. In: Tile PM, ed. Bailey and Scott’s Diagnostic Microbiology. 14th ed. St Louis, MO: Mosby Inc; 2017:1046-1054.

    • Search Google Scholar
    • Export Citation
  • 83.

    Wilcox ME, Chong CAKY, Stanbrook MB, et al.Does this patient have an exudative pleural effusion? The Rational Clinical Examination systematic review. JAMA. 2014;311(23):2422-2431.PubMed

    • Search Google Scholar
    • Export Citation
  • 84.

    Penn RL, Betts RF. Lower respiratory tract infections (including tuberculosis). In: Betts RF, Chapman SW, Penn RL, eds. A Practical Approach to Infectious Diseases. 5th ed. Philadelphia: Lippincott, Williams and Wilkins; 2003:295-371.

    • Search Google Scholar
    • Export Citation
  • 85.

    Parta M. Pleural effusion and empyema. In: Bennett JE, Dolin R, Blaser MJ, eds. Principles and Practice of Infectious Diseases. 9th ed. Philadelphia, PA: Elsevier, Inc.; 2020:914-925.

    • Search Google Scholar
    • Export Citation
  • 86.

    Brannan SR, Jerrard DA. Synovial fluid analysis. J Emerg Med. 2006;30(3):331-339.PubMed

  • 87.

    Ohl CA. Infectious arthritis of native joints. In: Bennett JE, Dolin R, Blaser MJ, eds. Principles and Practice of Infectious Diseases. 9th ed. Philadelphia, PA: Elsevier, Inc.; 2020:1400-1417.

    • Search Google Scholar
    • Export Citation
  • 88.

    Bergman SJ, Armstrong EP. Bone and joint infections. In: DiPiro JT, Yee GC, Posey L, et al., eds. Pharmacotherapy: A Pathophysiologic Approach, 11th ed. New York, NY: McGraw-Hill; 2020. https://accesspharmacy.mhmedical.com/content.aspx?bookid=2577&sectionid=219306547. Accessed June 29, 2020.

    • Search Google Scholar
    • Export Citation
  • 89.

    Bush LM, Levison ME. Peritonitis and intraperitoneal abscesses. In: Bennett JE, Dolin R, Blaser MJ, eds. Principles and Practice of Infectious Diseases. 9th ed. Philadelphia, PA: Elsevier, Inc.; 2020:1009-1036.

    • Search Google Scholar
    • Export Citation
  • 90.

    Riggio O, Angeloni S. Ascitic fluid analysis for diagnosis and monitoring of spontaneous bacterial peritonitis. World J Gastroenterol. 2009;15(31):3845-3850.PubMed

    • Search Google Scholar
    • Export Citation
  • 91.

    Dayer E, Dayer JM, Roux-Lombard P. Primer: the practical use of biological markers of rheumatic and systemic inflammatory diseases. Nat Clin Pract Rheumatol. 2007;3(9):512-520.PubMed

    • Search Google Scholar
    • Export Citation
  • 92.

    Markanday A. Acute phase reactants in infections: evidence-based review and a guide for clinicians. Open Forum Infect Dis. 2015;2(3):ofv098.PubMed

    • Search Google Scholar
    • Export Citation
  • 93.

    Young B, Gleeson M, Cripps AW. C-reactive protein: a critical review. Pathology. 1991;23(2):118-124.PubMed

  • 94.

    Hogevik H, Olaison L, Andersson R, et al.C-reactive protein is more sensitive than erythrocyte sedimentation rate for diagnosis of infective endocarditis. Infection. 1997;25(2):82-85.PubMed

    • Search Google Scholar
    • Export Citation
  • 95.

    Olaison L, Hogevik H, Alestig K. Fever, C-reactive protein, and other acute-phase reactants during treatment of infective endocarditis. Arch Intern Med. 1997;157(8):885-892.PubMed

    • Search Google Scholar
    • Export Citation
  • 96.

    Heiro M, Helenius H, Sundell J, et al.Utility of serum C-reactive protein in assessing the outcome of infective endocarditis. Eur Heart J. 2005;26(18):1873-1881.PubMed

    • Search Google Scholar
    • Export Citation
  • 97.

    Davies J. Procalcitonin. J Clin Pathol. 2015;68:675-679.PubMed

  • 98.

    Schuetz P, Albrich W, Mueller B. Procalcitonin for diagnosis of infection and guide to antibiotic decisions: past, present and future. BMC Med. 2011;9:107.PubMed

    • Search Google Scholar
    • Export Citation
  • 99.

    Gilbert DN. Use of plasma procalcitonin levels as an adjunct to clinical microbiology. J Clin Microbiol. 2010;48(7):2325-2329.PubMed

  • 100.

    Vincent JL, Van Nuffelen M, Lelubre C. Host response biomarkers in sepsis: the role of procalcitonin. In: Mancini N, ed. Sepsis: Diagnostic Methods and Protocols. New York, NY: Humana Press; 2015:213-224.

    • Search Google Scholar
    • Export Citation
  • 101.

    Azzini AM, Dorizzi RM, Sette P, et al.A 2020 review on the role of procalcitonin in different clinical settings: an update conducted with the tools of the evidence based laboratory medicine. Ann Transl Med. 2020;8(9):610.PubMed

    • Search Google Scholar
    • Export Citation