What is Dr. Zubair Khalid's research focus?

Dr. Zubair Khalid specializes in molecular virology, mRNA vaccine development, and computational biology, with a focus on avian pathogens like IBDV and Avian Reovirus.

Where is Dr. Zubair Khalid currently working?

Dr. Zubair Khalid is a Postdoctoral Research Associate at the University of Maryland (UMD), specifically within the Department of Animal and Avian Sciences.

How to Calculate the Number of Bacteria in a Sample Using the Drop Plate Method

Q: 1. Why is the countable range for drops smaller than for spread plates?

The countable range for drops (3–30 colonies per drop) is narrower than for spread plates (30–300 colonies per plate) because the small volume limits the number of colonies that can be resolved without overlap. In a 10 µL drop, colonies are concentrated in a small area, so even 30 colonies may be difficult to count accurately. The statistical reliability also decreases at very low counts (below 3 colonies) because the sampling error becomes large relative to the count.

Q: 2. Can I use the drop plate method for anaerobic bacteria?

Yes, but you must incubate plates in an anaerobic chamber or jar with an appropriate gas-generating system. The drops must be placed on prereduced agar medium, and the plates should be transferred to anaerobic conditions immediately after the drops have absorbed. The same calculation formula applies, but ensure that the countable range is validated for the specific organism under anaerobic conditions.

Q: 3. How do I handle samples with very high or very low bacterial concentrations?

For high concentrations, prepare additional 10-fold dilutions until you reach a dilution that yields 3–30 colonies per drop. For low concentrations, use the undiluted sample and plate multiple drops (e.g., 10 drops of 10 µL each) to increase the total volume plated. If even the undiluted sample yields no colonies, report as "less than 100 CFU/mL" (for 10 µL drops) or use membrane filtration for greater sensitivity.

Q: 4. What is the difference between the drop plate method and the Miles and Misra method?

The terms are often used interchangeably. The original Miles and Misra method (1938) used a calibrated dropping pipette to deliver 0.02 mL (20 µL) drops onto agar plates. Modern adaptations use micropipettes to deliver 10–20 µL drops. The principle, calculation, and countable range are the same. Some protocols refer specifically to the Miles and Misra method when using 20 µL drops and a specific grid pattern, but the core technique is identical.

Detailed view of a microscope in a laboratory used in scientific research — Photo by indra projects on Pexels.

The drop plate method (also known as the Miles and Misra method) is a bacterial enumeration technique where small, calibrated volumes of serial dilutions are deposited as discrete drops onto the surface of an agar plate, allowed to dry, and incubated to produce countable colonies. This method is particularly useful when sample volume is limited, when rapid screening of multiple dilutions is needed, or when working with non-spreading organisms that form discrete colonies. The calculation converts colony counts from countable drops into colony-forming units per milliliter (CFU/mL) using a formula that accounts for the volume plated and the dilution factor.

At a Glance

Aspect	Detail
Method name	Drop plate method (Miles and Misra method)
Purpose	Enumeration of viable bacteria in a liquid sample
Sample volume per drop	Typically 10–20 µL (standardized by pipette)
Plating pattern	3–5 drops per dilution, placed in sectors on one plate
Countable range	3–30 colonies per drop (or 5–50 per drop, depending on local SOP)
Calculation formula	CFU/mL = (Average colonies per drop) × (1/volume per drop in mL) × (1/dilution factor)
Key advantage	Multiple dilutions on a single plate, conserving materials
Key limitation	Requires non-swarming organisms; drops may coalesce if too large
Biosafety level	BSL-1 for non-pathogenic organisms; follow institutional risk assessment

Scientific Principle

The drop plate method relies on the same fundamental principle as other viable count techniques: each viable bacterial cell, when deposited on a suitable solid growth medium, will divide to form a single visible colony. The method assumes that one colony arises from one viable cell (or one colony-forming unit, CFU), which may represent a single cell or a clump of cells.

The key distinction of the drop plate method is the plating geometry. Instead of spreading a volume across the entire plate surface, small drops (typically 10–20 µL) are placed in a grid pattern. This allows multiple dilutions to be tested on a single plate, reducing the number of plates needed and the overall material cost. The method was originally described by Miles and Misra in 1938 and remains widely used in research and teaching laboratories.

The calculation corrects for the small volume plated and the dilution applied to the original sample. The countable range for drops is narrower than for spread plates because the small volume limits the number of colonies that can be resolved without overlapping. Most protocols accept 3–30 colonies per drop as the countable range, though some laboratories extend this to 5–50 colonies per drop for certain organisms.

Materials and Instrumentation Choices

Agar Plates

The choice of agar medium depends on the target organism. For general enumeration of non-fastidious bacteria, tryptic soy agar (TSA) or nutrient agar is appropriate. For selective enumeration, use media containing antibiotics or specific substrates. Plates should be dried before use to ensure drops are absorbed quickly without spreading. Pre-dry plates by leaving them open in a biosafety cabinet for 15–30 minutes, or use commercially prepared plates that are sufficiently dry.

Pipettes and Tips

Accurate volume delivery is critical. Use calibrated micropipettes capable of delivering 10–20 µL with precision. Positive displacement pipettes are recommended for viscous samples or when working with small volumes. Always use sterile, aerosol-resistant tips to prevent cross-contamination.

Dilution Tubes and Diluents

Use sterile dilution blanks containing phosphate-buffered saline (PBS), 0.85% saline, or 0.1% peptone water. The diluent should maintain cell viability without supporting growth. Prepare serial dilutions in a logical sequence (e.g., 10⁻¹ through 10⁻⁷) based on the expected bacterial concentration.

Incubator

Standard incubation conditions (35–37°C for mesophilic bacteria) apply. Ensure the incubator is calibrated and maintains stable temperature. Incubation time depends on the organism; most bacteria form visible colonies within 18–24 hours.

Colony Counter

A manual colony counter with a magnifying lens and grid is sufficient. Automated colony counters can be used but must be validated for drop plate patterns. Deep learning-based colony classification systems have shown high accuracy for automated counting [2], but these systems require training on the specific plate format and organism.

Controls

Positive Control

Include a reference strain with known growth characteristics to verify that the medium and incubation conditions support growth. For BSL-1 work, Escherichia coli K-12 or Bacillus subtilis are suitable.

Negative Control

Plate sterile diluent alone to confirm that the diluent and pipetting technique do not introduce contaminants. No colonies should appear on negative control drops.

Dilution Control

Plate the same dilution in duplicate or triplicate to assess reproducibility. The coefficient of variation between replicate drops should be less than 30% for reliable results.

Volume Control

Weigh the delivered volume periodically to verify pipette calibration. For a 10 µL pipette set to deliver 10 µL, the actual delivered volume should be within 1% of the set value.

Conceptual Workflow

Step 1: Prepare Serial Dilutions

Label sterile dilution tubes with the dilution factor (e.g., 10⁻¹, 10⁻², 10⁻³). Add 900 µL of sterile diluent to each tube. Transfer 100 µL of the original sample to the 10⁻¹ tube and vortex thoroughly. Using a fresh tip, transfer 100 µL from the 10⁻¹ tube to the 10⁻² tube, and continue through the desired dilution series. Vortex each tube for at least 5 seconds before proceeding.

Step 2: Plate the Drops

Divide the agar plate into sectors (typically 4–6 sectors) by marking the bottom of the plate with a permanent marker. Label each sector with the dilution factor. Using a calibrated micropipette, deposit 10 µL (or your chosen volume) of each dilution as a single drop in the center of its sector. For each dilution, deposit 3–5 replicate drops in separate positions within the sector. Work quickly to prevent drops from drying before plating.

Step 3: Allow Drops to Absorb

Leave the plates open in a biosafety cabinet for 15–30 minutes until the drops have fully absorbed into the agar. Do not move the plates during this time, as drops may coalesce. If drops are not absorbing, the agar may be too wet; pre-dry plates more thoroughly in future experiments.

Step 4: Incubate

Invert the plates and incubate at the appropriate temperature for 18–24 hours. For slow-growing organisms, extend incubation time as needed. Check plates at 24 hours and again at 48 hours if necessary.

Step 5: Count Colonies

Select the dilution sector where drops contain 3–30 colonies each. Count all colonies in each of the 3–5 replicate drops. Record the counts for each drop separately. If no sector meets the countable range, select the sector closest to the range and note the limitation.

Step 6: Calculate CFU/mL

Use the following formula:

CFU/mL = (Average number of colonies per drop) × (1 / Volume per drop in mL) × (1 / Dilution factor)

For example, if you plated 10 µL of the 10⁻⁵ dilution and counted an average of 20 colonies per drop:

CFU/mL = 20 × (1 / 0.01 mL) × (1 / 10⁻⁵) = 20 × 100 × 100,000 = 2.0 × 10⁸ CFU/mL

Step 7: Report Results

Report the result as CFU/mL with the dilution used and the volume plated. Include the countable range and the number of replicates. For example: "2.0 × 10⁸ CFU/mL (10 µL drops of 10⁻⁵ dilution, 5 replicates, countable range 3–30 colonies per drop)."

Quality Checks

Verify Pipette Calibration

Before each experiment, verify that the pipette delivers the correct volume. For a 10 µL pipette, dispense 10 µL onto a precision balance and record the weight. The weight in milligrams should equal the volume in microliters (assuming water density). Repeat three times and calculate the mean.

Check Drop Morphology

After absorption, drops should form circular, discrete zones. If drops have spread irregularly or merged, the agar was too wet or the drops were placed too close together. Discard such plates and repeat with properly dried plates.

Assess Colony Morphology

Colonies within a drop should be uniform in appearance. If colonies show mixed morphologies, the sample may contain multiple species, or contamination may have occurred. Record observations and consider repeating with selective media.

Evaluate Replicate Consistency

Calculate the coefficient of variation (CV) for replicate drops: CV = (Standard deviation / Mean) × 100%. A CV below 30% indicates acceptable reproducibility. Higher CV suggests pipetting errors, uneven mixing, or clumping of cells.

Result Interpretation

Acceptable Counts

The countable range of 3–30 colonies per drop is based on statistical reliability. Below 3 colonies, the sampling error is too high; above 30 colonies, colony overlap and counting errors increase. Some protocols extend the range to 5–50 colonies per drop for organisms that form very small colonies, but this should be validated for each organism.

Outlier Handling

If one replicate drop shows a count that is dramatically different from the others (e.g., 25, 28, and 2 colonies), investigate the outlier. Possible causes include a contaminated pipette tip, a drop that was not properly mixed, or a drop that was placed on a defective area of the agar. Exclude the outlier only if a clear technical error is identified; otherwise, include all counts.

Zero Counts

If no colonies appear in any dilution, the sample may contain fewer than 10 CFU/mL (the detection limit for a 10 µL drop of the undiluted sample). Report as "less than 10 CFU/mL" or repeat with a larger sample volume using membrane filtration.

Too Many Colonies

If all drops show >30 colonies, repeat the experiment with higher dilutions. If the highest dilution still shows >30 colonies, the sample concentration exceeds the method's capacity. Consider using the spread plate method with larger volumes for more accurate enumeration.

Troubleshooting

Observation	Likely Cause	Discriminating Check
Drops spread and merge on plate	Agar too wet	Pre-dry plates for 30–60 minutes before use
No colonies in any dilution	Sample too dilute, or cells non-viable	Plate undiluted sample; check viability with positive control
Colonies too numerous to count in all dilutions	Insufficient dilution series	Prepare additional 10-fold dilutions (e.g., 10⁻⁶, 10⁻⁷)
High variability between replicate drops	Incomplete mixing of dilution tubes	Vortex each dilution tube for 10 seconds before pipetting
Colonies appear only on plate edges	Drops placed too close to edge; edge effect	Place drops at least 1 cm from plate edge
Colonies are tiny and difficult to count	Incubation time too short or medium inadequate	Extend incubation; verify medium supports growth
Irregular colony shapes within a drop	Mixed culture or contamination	Streak a colony for isolation; perform Gram stain
Drops do not absorb into agar	Agar surface too dry or too wet	Adjust pre-drying time; use freshly poured plates

Limitations

Organism Restrictions

The drop plate method is not suitable for swarming organisms such as Proteus species or Serratia marcescens, which spread across the agar surface and obscure individual colonies. For these organisms, use the pour plate method or add swarming inhibitors to the medium.

Volume Constraints

The small volume per drop (10–20 µL) limits the detection sensitivity. For samples with low bacterial concentrations (<10³ CFU/mL), the drop plate method may not yield countable drops. Use membrane filtration or the spread plate method with larger volumes for such samples.

Clumping and Chains

Bacteria that form chains (e.g., Streptococcus species) or clumps will produce one colony from multiple cells, leading to underestimation of the true cell count. Sonication or vortexing with glass beads can help disperse clumps, but the result will still reflect CFU rather than individual cells.

Statistical Precision

The drop plate method has lower precision than the spread plate method because of the smaller volume plated. The confidence interval for a count of 20 colonies from a 10 µL drop is wider than for 200 colonies from a 100 µL spread plate. For applications requiring high precision, use the spread plate method with larger volumes.

Documentation

Laboratory Notebook Entry

Record the following information for each experiment:

Date and time of plating
Sample identification and source
Dilution series used (e.g., 10⁻¹ through 10⁻⁷)
Volume per drop (e.g., 10 µL)
Number of replicate drops per dilution
Agar medium type and lot number
Incubation temperature and duration
Raw colony counts for each drop
Calculated CFU/mL with dilution and volume correction
Any observations (e.g., colony morphology, contamination)

Data Reporting

When reporting results, include:

The method used (drop plate method)
The countable range applied (e.g., 3–30 colonies per drop)
The dilution and volume used for calculation
The number of replicate drops
The final CFU/mL value with appropriate significant figures
Any limitations or deviations from the standard protocol

Quality Records

Maintain records of:

Pipette calibration dates and results
Incubator temperature logs
Medium preparation and sterility testing
Positive and negative control results

Biosafety Considerations

The drop plate method is classified as a BSL-1 procedure when working with non-pathogenic organisms such as Escherichia coli K-12, Bacillus subtilis, or Rhodococcus species. Follow standard microbiological practices as outlined in the Biosafety in Microbiological and Biomedical Laboratories (BMBL) 6th Edition [6]:

Perform all work in a certified biosafety cabinet (BSC) to contain aerosols generated during pipetting and vortexing.
Use aerosol-resistant pipette tips to prevent contamination of the pipette barrel.
Decontaminate all waste (plates, tips, tubes) by autoclaving before disposal.
Wear appropriate personal protective equipment (PPE): lab coat, gloves, and eye protection.
Disinfect work surfaces before and after each session with 70% ethanol or 10% bleach.
Never pipette by mouth.
Wash hands thoroughly after handling cultures.

For work with recombinant or synthetic nucleic acid molecules, consult the NIH Guidelines for Research Involving Recombinant or Synthetic Nucleic Acid Molecules [7] and obtain institutional biosafety committee approval if required.

If the sample contains organisms that are not BSL-1, perform a risk assessment following institutional guidelines. The drop plate method may still be used at higher containment levels, but additional precautions (e.g., sealed plates, secondary containment) are necessary.

Frequently Asked Questions

1. Why is the countable range for drops smaller than for spread plates?