Antibiotic resistance is tested by exposing bacteria to different antibiotics in a lab and observing whether the bacteria survive, die, or keep growing. This process, called antimicrobial susceptibility testing (AST), is the foundation of how doctors choose which antibiotic to prescribe. There are several methods, ranging from simple agar plates to automated machines and genetic analysis, each with different strengths and turnaround times.
The Two Main Approaches
Resistance testing falls into two broad categories: phenotypic and genotypic. Phenotypic testing watches what the bacteria actually do when exposed to an antibiotic. You grow the organism in the presence of the drug and see if it survives. This remains the gold standard because it reflects real-world behavior. Genotypic testing, by contrast, looks at the bacteria’s DNA for known resistance genes. It’s faster, sometimes delivering results in under two hours compared to a full day or more for culture-based methods, but it has a significant limitation: finding a resistance gene doesn’t always mean the bacterium will behave as resistant. The gene may be present but not actively expressed, which can lead to a misleading result.
Most clinical labs rely on phenotypic methods as the primary tool and use molecular testing as a supplement, particularly when speed matters or the organism is difficult to grow in culture.
Disk Diffusion (Kirby-Bauer Test)
The disk diffusion test is one of the oldest and most widely used methods. A technician spreads a standardized amount of bacteria across an agar plate, then places small paper disks soaked in different antibiotics onto the surface. The plate goes into an incubator overnight, typically 16 to 18 hours. As the antibiotic diffuses outward from each disk, it creates a concentration gradient in the agar. If the bacteria are susceptible, a clear circle forms around the disk where nothing grew. If they’re resistant, bacteria grow right up to the edge.
The technician measures the diameter of each clear zone in millimeters, reading from the back of the plate held above a dark, nonreflecting surface under reflected light. Those measurements are then compared against published breakpoint tables to classify the bacteria as susceptible, intermediate, or resistant to each drug. The method is inexpensive and straightforward, which makes it practical for labs with limited resources, but it provides a qualitative result rather than a precise concentration value.
Minimum Inhibitory Concentration (MIC)
The MIC test answers a more specific question: what is the lowest concentration of an antibiotic that stops visible bacterial growth? To find out, a lab prepares a series of tubes or wells containing the same antibiotic at progressively higher concentrations, each created through serial dilution. The bacteria are added to every well at a standardized density, then incubated for 18 to 24 hours at body temperature. The technician examines each well afterward. The first concentration where no visible growth appears is the MIC value.
This number is clinically useful because it tells a doctor not just whether an antibiotic works, but how much is needed. A bacterium with a low MIC for a given drug is highly susceptible; one with an MIC just below the resistance threshold may still be treatable but requires careful dosing. Broth microdilution, which uses 96-well plates to test many antibiotics at once, is the reference standard for MIC determination.
Gradient Strip Testing
Gradient strips (commonly known by the brand name Etest) combine elements of disk diffusion and MIC testing. A plastic or paper strip carries a predefined exponential gradient of antibiotic along its length, with concentration values printed on the surface. When placed on an inoculated agar plate, the drug releases immediately into the medium. After incubation, bacterial growth forms an elliptical zone of inhibition around the strip. The MIC is read where the edge of the ellipse intersects the concentration scale on the strip.
This method gives you a quantitative MIC value without needing to set up a full serial dilution, making it a practical option when a lab needs precise numbers for a small number of antibiotics.
Automated Systems in Hospital Labs
High-volume clinical microbiology labs typically use automated platforms that standardize and speed up the entire process. The most common systems include the BD Phoenix, Vitek 2, and MicroScan. These instruments prepare bacterial suspensions, expose them to panels of antibiotics at multiple concentrations, and use optical sensors to detect growth over time. Built-in software then interprets results against current breakpoint standards and flags unusual resistance patterns.
Automated systems improve reproducibility and reduce human error compared to manual reading of plates, though they aren’t infallible. Studies comparing these platforms have found occasional errors in categorization, particularly with certain organism-drug combinations. Routine automated systems typically deliver results in 9 to 19 hours after the instrument receives the sample, though the total turnaround time from when a specimen arrives at the lab to when a result reaches the doctor can stretch to 22 to 45 hours because of the time spent on initial culture and identification steps.
Newer rapid AST technologies are compressing that timeline significantly. One system evaluated across multiple hospitals in 2025, called QuickMIC, produced results in an average of about 3 hours of instrument time, with a total lab turnaround of 10 to 11.5 hours. That difference matters most for serious infections like bloodstream infections, where every hour of delay in effective treatment affects outcomes.
Molecular and Genetic Testing
Instead of growing bacteria and watching their behavior, molecular methods look directly at their genetic code for known resistance markers. PCR-based tests can detect specific genes associated with resistance, such as genes that produce enzymes capable of breaking down certain antibiotics. These tests can be run directly on clinical samples like blood or urine, skipping the culture step entirely and returning results in as little as one to two hours.
Whole genome sequencing goes further, reading the organism’s entire DNA to identify every known resistance gene at once. This is especially valuable for slow-growing bacteria like tuberculosis, where traditional culture can take weeks. It also supports surveillance programs that track how resistance is spreading through a community or hospital.
The trade-off is that molecular tests can only find what they’re designed to look for. If a bacterium carries a novel or uncommon resistance mechanism not included in the test panel, it will be missed entirely. And because gene presence doesn’t always equal gene expression, a positive genetic result sometimes overpredicts resistance, potentially steering treatment away from an antibiotic that would have worked.
Mass Spectrometry for Resistance Detection
A technology originally used for rapid bacterial identification, called MALDI-TOF mass spectrometry, has been adapted to detect certain types of resistance. The approach works by incubating bacteria with an antibiotic and then using the instrument to detect whether the drug has been broken down. If the mass spectrum shows breakdown products of the antibiotic alongside a decrease in the intact drug, the bacterium is producing enzymes that destroy it. This method has proven particularly accurate for detecting bacteria that produce carbapenemases, enzymes that inactivate a critically important class of last-resort antibiotics. It can be performed directly from positive blood cultures and is fast enough to be practical in routine clinical lab workflows.
How Results Are Reported
Regardless of the method used, resistance test results are ultimately reported using three categories. “Susceptible” (S) means the antibiotic is expected to work at standard doses for the type of infection being treated. “Resistant” (R) means treatment with that antibiotic is likely to fail regardless of how it’s dosed. The middle category, labeled “I,” was traditionally called “intermediate” but has been redefined by the European Committee on Antimicrobial Susceptibility Testing (EUCAST) as “susceptible, increased exposure.” This means the antibiotic can still work, but only if the dose is increased or if the drug naturally concentrates at the infection site, such as urinary tract infections where certain antibiotics reach much higher levels in urine than in blood.
The cutoff values that define these categories are called breakpoints. Two organizations set them globally: EUCAST, used predominantly in Europe, and the Clinical and Laboratory Standards Institute (CLSI), used widely in the United States and other regions. These breakpoints are updated regularly as new resistance data emerges, which is why a bacterium classified as susceptible one year could be reclassified as intermediate or resistant the next if the thresholds shift.
What Determines Which Test Gets Used
The choice of method depends on the clinical situation, the organism involved, and the lab’s resources. Simple urinary tract infections might only need a disk diffusion test. Bloodstream infections in a critically ill patient call for the fastest available method, potentially combining molecular screening with automated phenotypic testing. Surveillance programs tracking resistance trends across a hospital system may use whole genome sequencing to map the spread of resistant strains.
In practice, many labs use a layered approach: rapid molecular tests to guide initial treatment decisions within hours, followed by definitive phenotypic results the next day that confirm or adjust the antibiotic choice. The goal across all methods is the same: match the right antibiotic to the specific bacteria causing the infection, at the right dose, as quickly as possible.