A serial dilution is a stepwise process where you repeatedly dilute a solution by the same factor, using the previous dilution as the starting point for the next one. Each step reduces the concentration by a fixed amount, letting you quickly span a wide range of concentrations from a single stock solution. The technique is used constantly in microbiology, chemistry, pharmacology, and molecular biology.
How Serial Dilutions Work
Instead of making each dilution independently from your original stock, you make them in a chain. You transfer a small volume from your stock into a tube of diluent, mix thoroughly, then transfer the same volume from that tube into the next one. Each transfer multiplies the previous dilution by the same factor. After just a few steps, you can reduce a concentration by thousands or even millions of times.
The math behind each individual step follows a simple relationship: the starting concentration times the starting volume equals the final concentration times the final volume. Written out, that’s C1 × V1 = C2 × V2. But the real power of serial dilutions is that the dilution factors multiply across steps. If each step is a 1:10 dilution, then after two steps you have 1:100, after three steps 1:1,000, and so on.
Choosing a Dilution Factor
The two most common series are tenfold (1:10) and twofold (1:2), and each suits different situations.
- Tenfold (1:10): You add 1 part sample to 9 parts diluent, making a total volume of 10. Each step drops the concentration by a factor of ten. This is the standard in microbiology when you need to reduce bacterial counts across many orders of magnitude for plating and colony counting. Three tubes give you 1:1,000; six tubes give you 1:1,000,000.
- Twofold (1:2): You add 1 part sample to 1 part diluent. Each step cuts the concentration in half. This is common in immunology, pharmacology, and any assay where you need finer resolution between concentrations, such as determining the minimum inhibitory concentration of an antibiotic or building a standard curve for an ELISA.
Your choice depends on the range you need to cover and how much detail you need between points. Tenfold dilutions cover enormous ranges quickly but leave large gaps between concentrations. Twofold dilutions give you more data points in a narrower range.
Step-by-Step: Making a Tenfold Series
Here’s the standard protocol for a 1:10 serial dilution, the version you’ll encounter most often in microbiology coursework and lab work.
1. Prepare your tubes. Line up the number of tubes you need (typically 5 to 8). Add 9 ml of sterile diluent to each one. Common diluents include sterile saline, phosphate-buffered saline, or sterile water, depending on what you’re diluting and what the downstream application requires. Label each tube with its dilution factor: 10⁻¹, 10⁻², 10⁻³, and so on.
2. Transfer from the stock. Using a sterile pipette, draw up 1 ml of your original sample and add it to the first tube (which contains 9 ml of diluent). You now have 1 ml of sample in 10 ml total, giving you a 1:10 dilution.
3. Mix thoroughly. Vortex the tube or invert it several times. Inadequate mixing is one of the most common sources of error. The sample needs to be evenly distributed before you pull from this tube for the next step.
4. Use a fresh pipette. Discard the pipette you just used and pick up a new sterile one. Reusing pipettes between steps introduces carryover that throws off your concentrations.
5. Transfer to the next tube. Draw 1 ml from the first dilution tube and add it to the second tube. This second tube now contains a 1:100 total dilution of the original sample (1/10 × 1/10 = 1/100).
6. Repeat. Continue mixing, changing pipettes, and transferring 1 ml to the next tube for as many steps as you need. Each tube is ten times more dilute than the one before it.
You can adjust volumes as long as you keep the ratio. For example, the American Society for Microbiology notes that adding 0.5 ml to 4.5 ml gives the same 1:10 dilution (0.5 divided by 5.0 equals 1/10). Smaller volumes save reagents when supplies are limited.
Calculating the Final Concentration
To find the concentration at any step, multiply the original concentration by the dilution factor raised to the number of steps. If you started with 1,000,000 bacteria per ml and performed three tenfold dilutions, the third tube contains 1,000,000 × (1/10)³ = 1,000 bacteria per ml.
You can also think of it as multiplying the dilution of each tube by the dilution of the tube before it. The ASM protocol illustrates this clearly: a 1:10 dilution in the first tube multiplied by a 1:10 dilution in the second tube gives a total dilution of 1:100 in the second tube. The third tube would be 1:1,000, and so on. This cumulative calculation is essential for back-calculating the original concentration of your sample after you count colonies or measure absorbance at the end.
Building a Standard Curve
Serial dilutions are the backbone of standard curves in assays like ELISA, qPCR, and spectrophotometry. You start with a known concentration of your target molecule (an antibody, a piece of DNA, a protein) and dilute it in a series to create a set of known reference points. You then plot the instrument’s response (color intensity, fluorescence, absorbance) against those known concentrations. When you run your unknown samples, you compare their signal to the curve to determine their concentration.
Before designing a dilution series for a standard curve, you need a rough estimate of where your unknown samples fall. The goal is to bracket the expected concentration so that your unknowns land somewhere in the middle of the curve, where the measurements are most reliable. You also need to decide on the range: what’s the lowest and highest concentration you want to test? For assays that span several orders of magnitude, tenfold dilutions plotted on a logarithmic scale work well. For assays with a narrower dynamic range, twofold or threefold dilutions give you better resolution.
How Errors Accumulate
The single most important thing to understand about serial dilutions is that every small error compounds. Because each step uses the output of the previous step as its input, a pipetting mistake in tube one propagates through every tube that follows.
Research published in the Journal of Computer-Aided Molecular Design modeled this precisely. Even with a robotic liquid handler rated at just 3% imprecision and 3 to 5% inaccuracy per transfer, the errors grew with each step. The coefficient of variation (a measure of how scattered your results are) increased monotonically across the series. By the final well of an 8-point dilution series, the cumulative bias reached nearly 50%, meaning the actual concentration was about half of what it should have been. Extend the series to 16 or 32 wells and the bias approached 100%, essentially rendering the final wells meaningless.
This has practical implications. If your experiment demands high accuracy at the far end of a long dilution series, you may be better off preparing those concentrations independently from the stock rather than relying on a serial chain. For most routine lab work with 5 to 8 dilution steps, careful pipetting keeps the error manageable.
Tips for Better Accuracy
Most dilution errors come down to three things: pipetting, mixing, and contamination. Here’s how to minimize each one.
Pipetting is the foundation. Use calibrated pipettes and check that you’re dispensing the correct volume. Pre-wet the pipette tip by aspirating and dispensing the liquid once before taking the actual measurement, especially with viscous solutions. Always pipette slowly and consistently. Hold the pipette vertically and touch the tip to the inside wall of the receiving tube to ensure complete delivery.
Mixing matters more than most people realize. The ASM protocol emphasizes that accuracy in quantitation depends on adequate agitation of dilution tubes. After adding your sample to the diluent, vortex or vigorously invert the tube at least 5 to 10 times. If you transfer from an insufficiently mixed tube, you’ll pull a sample that doesn’t represent the true concentration, and that error carries forward through every subsequent tube.
Use a fresh, sterile pipette or tip for every transfer. This prevents carrying extra liquid between tubes, which would change the dilution factor, and eliminates cross-contamination. In microbiology work, sterile technique throughout the process is non-negotiable: any contaminant introduced at an early step will be amplified through the remaining tubes and show up as false colonies on your plates.
Colony Counting in Microbiology
One of the most common uses for serial dilutions is determining how many bacteria are in a sample. You dilute the sample in a tenfold series, plate a known volume from each tube onto agar, incubate overnight, and count the colonies that grow. The plates you actually count are the ones with a manageable number of colonies, typically between 30 and 300. Fewer than 30 and random variation makes the count unreliable. More than 300 and the colonies merge together and become impossible to distinguish.
To calculate the original concentration, you take the colony count, divide by the volume you plated, and divide by the dilution factor of that tube. If you counted 150 colonies on a plate from the 10⁻⁵ dilution tube, and you plated 0.1 ml, the original sample contained 150 ÷ 0.1 ÷ 10⁻⁵ = 15,000,000 (or 1.5 × 10⁷) colony-forming units per ml.