It sounds simple, and the underlying principle is. But serial dilution done incorrectly produces results that are wrong by orders of magnitude, and the errors often come from the same small set of mistakes that are straightforward to avoid once you know what they are. This page gives you the full procedure, the calculation with worked examples, the common mistakes and why they matter, and the practical decisions around choosing volumes, countable ranges, and plate types.
Why We Dilute: The Problem of Too Many Bacteria
A typical overnight liquid culture of E. coli contains approximately 10^9 CFU/mL. A contaminated food sample might contain anywhere from 10^2 to 10^8 CFU/g depending on the food and the degree of contamination. If you plate 0.1 mL of either of these samples directly onto agar, you will get a plate with so many colonies they merge into a continuous lawn. You cannot count them. You cannot distinguish individual colonies. The result is reported as Too Many To Count (TMTC) and is useless for quantification.
Serial dilution reduces the concentration stepwise, by a fixed factor at each step, until the expected concentration at some dilution step falls within the countable range. The countable range for a standard spread plate is 30 to 300 colonies. A plate with fewer than 30 colonies is Too Few To Count (TFTC) with adequate statistical confidence. A plate with more than 300 colonies is Too Many To Count. The goal of the dilution series is to have at least one dilution step produce a plate within this countable range.
The 10-Fold (1:10) Dilution Series
The standard serial dilution uses 10-fold steps. At each step, 1 part of the previous tube is added to 9 parts of diluent (water, physiological saline at 0.9 per cent NaCl, or phosphate buffer). This produces a 1:10 dilution at each step. After six steps, the concentration has been reduced from the original by a factor of 10^6 (one million-fold).
The dilution factor at each step is the inverse of the fraction of original sample present: a 1:10 dilution has a dilution factor of 10^-1. After six 1:10 steps, the dilution factor is 10^-6.
Procedure in detail: Prepare 6 labelled tubes, each containing exactly 9.0 mL of diluent (use a pipette or volumetric device, not a graduated cylinder, for accuracy). Add 1.0 mL of the original sample to Tube 1. Mix thoroughly by vortexing for 5 to 10 seconds. This is critical: inadequate mixing means cells from the previous tube are not uniformly distributed in the new tube, causing inaccurate results. Using a fresh pipette tip, transfer 1.0 mL from Tube 1 to Tube 2. Mix. Transfer 1.0 mL from Tube 2 to Tube 3. Mix. Continue through all six tubes. Each tube now contains a concentration 10 times lower than the previous tube.
From each tube (or from selected tubes based on expected concentration), plate 0.1 mL onto an agar plate. Note: plating 0.1 mL means the total dilution factor on the plate is 10 times lower than the dilution factor of the tube. A plate from the 10^-4 tube plated at 0.1 mL represents a dilution of 10^-5 on the plate.
Calculating CFU/mL
After incubation, count the colonies on plates within the countable range (30 to 300 colonies). Apply the formula:
CFU/mL = (Number of colonies counted) / (Volume plated in mL) x (Dilution factor of tube)
Equivalently: CFU/mL = (Number of colonies counted) / (Volume plated in mL x dilution of tube used)
Worked example: 145 colonies counted on the plate from the 10^-5 tube (labelled 10^-5 because 1.0 mL was transferred five times from original). Volume plated was 0.1 mL.
CFU/mL = 145 / (0.1 x 10^-5) = 145 / 10^-6 = 145 x 10^6 = 1.45 x 10^8 CFU/mL.
If results are obtained from two dilutions within the countable range (for example, 230 colonies from the 10^-5 plate and 41 colonies from the 10^-6 plate), the CFU/mL from both plates is calculated and the results are averaged if they agree within a factor of 2.
Choosing the Right Volume and Dilution Scheme
Standard protocol uses 1.0 mL transferred into 9.0 mL diluent at each step (1:10 dilution). For greater flexibility, you can use different volumes. Transferring 0.1 mL into 0.9 mL diluent also gives a 1:10 dilution but uses smaller volumes: useful when the sample is limited. Transferring 1.0 mL into 99 mL gives a 1:100 dilution in one step, useful for jumping quickly to very low concentrations.
Some applications use 1:2 dilutions (semi-log dilutions) when you need finer resolution around a specific concentration range: useful in MIC testing, dose-response experiments, and growth inhibition assays.
Spread Plate vs Pour Plate
The spread plate method is the standard for aerobic viable counts. A measured volume (0.1 mL is typical) is spread evenly across the surface of a pre-poured, solidified agar plate using a sterile spreader (glass or plastic L-shaped cell spreader or glass beads). Colonies grow on the agar surface and are easy to count and to pick for further work.
The pour plate method incorporates the sample into molten agar before pouring. A measured volume of the diluted sample is added to a tube of molten agar (held at 45 to 48 degrees Celsius to keep it liquid without killing the bacteria), mixed, and poured into a petri dish to solidify. Colonies grow both on the agar surface and within the agar. The pour plate allows plating of larger volumes (up to 1 mL compared to 0.1 mL for spread plate) and thus can detect lower concentrations. However, it requires careful temperature control of the molten agar, organisms must tolerate brief exposure to warm agar, and colonies within the agar can be harder to count and pick.
Common Mistakes in Serial Dilution
Not vortex-mixing between transfers: The single most common and most consequential error. If the previous tube is not thoroughly mixed before transferring the aliquot to the next tube, the concentration in the transfer volume does not represent the average concentration in the tube. Results become unpredictably inaccurate.
Using the same pipette tip across multiple transfers: Using the same tip from tube to tube carries the previous tube's concentration forward and prevents accurate dilution. Change tips between every dilution step.
Inaccurate volumes: Using a graduated cylinder or measuring cylinder for small volumes introduces significant volume errors. Use calibrated micropipettes for all volumes below 2 mL. Confirm calibration is current.
Letting molten agar cool before adding sample: In pour plate method, agar that has cooled below 44 degrees Celsius begins to solidify. Adding the sample to partially gelled agar produces uneven distribution and inaccurate colony counts.
Counting outside the countable range: Counting a plate with 320 colonies or 12 colonies introduces statistical error. If no plate from the dilution series falls within 30 to 300, note the results as approximate and consider re-running with a modified dilution range.
Not recording the plating volume: The calculation requires the exact volume plated. If 0.1 mL was intended but 0.12 mL was accidentally plated, the calculation will overestimate the concentration. Record all volumes at the time of plating.
Frequently Asked Questions
What is a serial dilution?
A serial dilution is a stepwise dilution of a sample, where each step reduces the concentration by a fixed factor. In microbiology, 10-fold (1:10) dilutions are standard. The series continues until the expected concentration at some step falls within the countable range for agar plating (30 to 300 colonies per plate for spread plates).
What is the countable range for microbiology plate counts?
The countable range for a spread plate is 30 to 300 colonies. Below 30 colonies the statistical confidence is too low (a small error in counting or an atypical distribution of colonies produces a large proportional error in the CFU/mL calculation). Above 300 colonies, individual colonies begin to merge and accurate counting becomes impossible.
How do you calculate CFU/mL from a plate count?
CFU/mL = Colony count / (Volume plated in mL x Dilution factor of the tube plated). For example, 156 colonies from a 10^-4 tube plated at 0.1 mL: CFU/mL = 156 / (0.1 x 10^-4) = 156 / 10^-5 = 1.56 x 10^7 CFU/mL.
Why must you change pipette tips between dilution steps?
Using the same pipette tip between dilution tubes carries the concentration from the previous tube forward, preventing accurate stepwise dilution. Each new step must start fresh with a clean, sterile tip to ensure the transferred volume accurately represents only the material from the previous tube at its correctly diluted concentration.
Why is vortex-mixing between transfers important?
Bacteria do not stay uniformly suspended in liquid: they settle, clump, and distribute unevenly. Thorough vortexing before each transfer ensures the aliquot removed represents the average concentration in the tube rather than a concentrated or dilute pocket. Inadequate mixing is one of the most common sources of serial dilution error.
What is TMTC and TFTC?
TMTC stands for "too many to count" and describes plates where colonies are so numerous they overlap and cannot be counted individually, generally more than 300 colonies per spread plate. TFTC stands for "too few to count" and describes plates with fewer than 30 colonies where the statistical reliability of the count is poor. Both results indicate the dilution factor chosen for that plate was not optimal and the dilution series should be extended or repeated with adjusted plating.
What diluent is used for serial dilutions?
Common diluents include 0.9 per cent physiological saline, phosphate-buffered saline (PBS), and 0.1 per cent peptone water. The choice depends on the application: peptone water is preferred for food microbiology because it provides a slight nutrient buffer that maintains viability of stressed cells during dilution. For pharmaceutical and environmental testing, specific diluents may be specified in the relevant pharmacopoeia or regulatory method.
What is the difference between a spread plate and a pour plate?
A spread plate is made by spreading a fixed volume (typically 0.1 mL) of diluted sample across the surface of pre-poured solidified agar. A pour plate mixes the diluted sample directly with molten agar, which is then poured and allowed to solidify. Spread plates allow larger colonies that are easier to count and pick. Pour plates allow larger plating volumes (useful for detecting very low concentrations) but require careful temperature control and produce subsurface colonies that are harder to pick.
How do you handle results from two plates within the countable range?
If two consecutive dilutions both produce plates within the 30 to 300 range (which sometimes happens at the boundary of the ideal dilution range), calculate CFU/mL from both plates independently. If the two results agree within a factor of 2, calculate the average and report it. If they differ by more than a factor of 2, investigate why: likely causes include pipetting error, inadequate mixing of a dilution step, or an inconsistently labelled tube.
What volume should be plated for a standard aerobic viable count?
For spread plates, 0.1 mL is standard. Larger volumes can be plated (up to 0.5 mL on a standard 90 mm plate) but require longer spreading time and care to avoid edge flooding. For pour plates, up to 1.0 mL of the diluted sample can be mixed into the molten agar before pouring, allowing detection of organisms present at concentrations as low as 1 CFU/mL when the 10^-1 or 10^0 dilution is plated.