Making an agarose gel is one of the most straightforward procedures in molecular biology: you dissolve agarose powder in a heated buffer, pour it into a casting tray with a comb, and let it cool until it solidifies into a translucent slab. The whole process takes roughly 30 to 45 minutes from start to finish. But the choices you make along the way, particularly the agarose concentration, the buffer, and the staining method, determine whether you get crisp, well-resolved bands or a blurry mess. Getting those details right matters far more than the pouring itself.
How Agarose Concentration Controls What You Can Separate
When agarose cools, its polymer chains associate and form a network of bundles with pores throughout the gel. The size of those pores is what separates DNA fragments: smaller fragments slip through easily, while larger ones get held up longer and migrate more slowly toward the positive electrode.1PubMed Central. Agarose gel electrophoresis for the separation of DNA fragments The concentration of agarose you use directly sets those pore sizes. A dilute gel (say, 0.5–0.8%) has large pores and works well for separating big DNA fragments in the range of several thousand base pairs and up. A denser gel (1.5–2%) has smaller pores and gives better resolution for fragments under a few hundred base pairs.
The relationship between concentration and pore size follows a predictable pattern: as you increase the agarose percentage, pore size shrinks and the distribution of pore sizes becomes narrower, meaning more uniform sieving.2PubMed. Pore size of agarose gels by atomic force microscopy For most routine cloning and PCR checks, a 1% gel is the default. It resolves fragments from roughly 200 bp to about 6,000 bp reasonably well and is easy to handle. If you are trying to distinguish fragments that are very close in size, you may need to go higher. Researchers have pushed agarose concentrations to 6–8% to resolve DNA fragments differing by as little as 2–5 base pairs in the 50–200 bp range, though this requires some buffer adjustments.3PubMed. A method for high-concentration agarose gel preparation and its application in high-resolution separation of low-molecular-weight nucleic acids and proteins For everyday lab work, though, you will rarely go beyond 2%.
A good rule of thumb: if you already know the approximate sizes of the fragments you expect, pick the concentration that puts your target size range in the middle of the gel’s useful separation window. Running a digest you have never seen before? Start with 1% and adjust from there.
TAE or TBE: Choosing a Buffer
You do not dissolve agarose in water. You use an electrophoresis buffer, and the two standard choices are TAE (Tris-acetate-EDTA) and TBE (Tris-borate-EDTA). Both maintain a stable pH during the run so your DNA stays negatively charged and migrates toward the positive electrode. For fragments ranging from several hundred to several thousand base pairs, both buffers deliver good resolution, with TBE giving slightly sharper bands for smaller fragments and TAE performing a bit better on larger ones.4PubMed Central. Modification of gel architecture and TBE/TAE buffer composition to minimize heating during agarose gel electrophoresis
Beyond resolution differences, there are practical reasons to favor one over the other. TAE is the go-to choice if you plan to cut a band out of the gel and recover the DNA for downstream work like ligation. Borate ions in TBE can inhibit enzymes used in cloning reactions, so TAE avoids that headache. TBE, on the other hand, has higher buffering capacity and is less prone to overheating during long runs at higher voltages, which makes it a better pick for extended electrophoresis or when you are running many gels in the same tank without changing the buffer.
Both buffers are typically made as concentrated stock solutions (often 10× or 50× for TAE, 5× or 10× for TBE) and diluted to 1× working concentration before use. The same buffer concentration used in the gel must also fill the electrophoresis tank. Mixing TAE gel with TBE running buffer, or vice versa, creates an ion mismatch that distorts migration and ruins resolution.
Step-by-Step Procedure
Here is how to actually make the gel, assuming a standard mini-gel setup and a 1% concentration. Adjust the agarose weight proportionally for other percentages.
- Weigh the agarose: For a 1% gel, measure 1 gram of agarose powder per 100 mL of buffer. Scale to your casting tray volume; most mini-gel trays need 30–50 mL, so you would use 0.3–0.5 g.
- Mix with buffer: Add the agarose to 1× TAE or TBE in an Erlenmeyer flask or bottle. Use a container at least twice the volume of your solution, because the liquid will boil up.
- Heat to dissolve: Microwave in short bursts (20–30 seconds each), swirling gently between intervals. Watch for boiling over. The solution is ready when it is completely clear with no visible particles or swirls. Undissolved agarose granules will create uneven pore sizes and distorted bands.
- Cool before pouring: Let the solution cool to about 55–60°C. You should be able to hold the flask comfortably, though it will still feel quite warm. If you are adding a pre-cast DNA stain, this is when you add it. Pouring too hot can warp the casting tray or crack the gel apparatus.
- Pour into the tray: Set the comb in the casting tray, then slowly pour the molten agarose. Tip the flask to one side of the tray and let the gel flow evenly. If you see air bubbles, push them to the edge with a pipette tip before the gel starts to set.
- Let it solidify: Leave the gel undisturbed at room temperature for 20–30 minutes. It turns from clear to slightly opaque when it has set. You can speed this up by placing the tray at 4°C for 10–15 minutes, though this is rarely necessary.
- Remove the comb: Pull the comb straight up gently. Yanking it at an angle can tear the wells. If the comb sticks, adding a small amount of running buffer on top of the gel before pulling usually loosens it.
- Submerge in buffer: Place the gel in the electrophoresis tank and add enough running buffer (the same buffer you used to make the gel) to cover the gel surface by 2–3 mm.
After loading your samples with loading dye into the wells, you are ready to run the gel. For a mini-gel, 80–120 volts for 30–60 minutes is a typical starting point. Lower voltage gives sharper resolution but takes longer; higher voltage is faster but generates more heat and can blur bands.
Staining and Seeing Your DNA
DNA is invisible on its own in the gel. You need a fluorescent dye to see it. The classic choice is ethidium bromide, which intercalates between DNA base pairs and fluoresces bright orange under UV light. It works well and is sensitive, but ethidium bromide is a known mutagen and requires careful handling and hazardous waste disposal.5PubMed. Comparative analysis of the DNA staining efficiencies of different fluorescent dyes in preparative agarose gel electrophoresis Many teaching labs and even research labs have shifted to safer alternatives like SYBR Gold and SYBR Green I, which are sensitive enough for most purposes and considerably less toxic. SYBR Gold is particularly good at picking up short fragments, while both dyes can detect 50 bp bands when given enough exposure time.5PubMed. Comparative analysis of the DNA staining efficiencies of different fluorescent dyes in preparative agarose gel electrophoresis One caveat: both SYBR dyes can alter how DNA migrates through the gel, which slightly shifts your size estimates compared to ethidium bromide. If precise sizing matters, run your ladder and samples with the same stain conditions.
You have three basic approaches to staining. You can add the dye directly to the molten agarose before pouring (pre-casting), soak the finished gel in a dye solution after the run (post-staining), or add the dye to the running buffer. Post-staining tends to produce the most sensitive and accurate results for band sizing, but it adds 20–50 minutes of soaking and washing time and uses more stain solution.6bioRxiv. A comparison of DNA stains and staining methods for Agarose Gel Electrophoresis Pre-casting is the most convenient for routine work: you add the dye once and the gel is ready to image the moment electrophoresis finishes. The trade-off is slightly lower sensitivity and the potential for dye-related mobility shifts. For quick-and-dirty PCR checks, pre-casting is perfectly fine. For publication-quality images where band sizes need to be accurate, post-staining is worth the extra time.
Common Problems and How to Fix Them
Smiling bands, where bands curve upward at the edges of the gel, almost always come from overheating. Running the gel at too high a voltage, using old or exhausted buffer, or letting the buffer level drop too low all contribute to uneven heat distribution. Lower the voltage, use fresh buffer, and make sure the gel is fully submerged.
Fuzzy or smeared bands have several possible causes. If every lane is smeared, the DNA may be degraded, the gel may have been loaded with too much DNA, or the gel itself may have set unevenly. If the smearing appears only in lanes with PCR products amplified from a complex template like environmental DNA, the explanation can be different: imperfect strand pairing between heterogeneous amplicon sequences creates double-stranded structures that migrate at unexpected rates, producing a smear even though the amplified DNA is all the correct size. Running the same sample on a denaturing alkaline gel can confirm whether the smear comes from size variation or structural artifacts.7PubMed. Band smearing of PCR amplified bacterial 16S rRNA genes: dependence on initial PCR target diversity
Wells that tear or collapse when loading samples usually mean the gel is too dilute for the comb teeth, or the comb was removed carelessly. Using a slightly higher agarose concentration (bumping from 0.8% to 1%) makes the gel more structurally sound. Loading buffer that is too dense can also punch through a fragile well bottom.
No bands at all? Check the obvious: did you remember to add loading dye, did you load the samples into the wells on the correct (cathode) end, and is the power supply actually delivering current? If the tracking dye ran in the right direction and the expected band still does not appear, the problem is likely upstream in the reaction that generated the DNA, not in the gel itself.
Low-Melting-Point Agarose for Gel Extraction
Standard agarose melts at around 85–95°C and re-solidifies near 35–40°C. That high melting point is a problem when you want to cut a band out of the gel and recover the DNA, because heating to melt the gel can damage your DNA or denature enzymes you plan to use. Low-melting-point (LMP) agarose is chemically modified to gel at lower temperatures, typically around 28–30°C, and melts at roughly 63–65°C.8International Journal of Biological Macromolecules. Oxyalkylation modification as a promising method for preparing low-melting-point agarose This means you can melt the gel slice at a gentler temperature before purifying the DNA.
LMP agarose costs more than standard agarose and produces softer, more fragile gels that can be tricky to handle. If you are planning to extract DNA from a gel, you can also use standard 1% agarose and dissolve the gel slice with chaotropic salts (the approach used by most commercial gel-extraction kits). At 1% concentration, gel slices dissolve readily, and the recovered DNA works well for downstream applications like ligation and transformation.9PubMed Central. An affordable and simple method for DNA extraction from agarose suitable for downstream applications Higher agarose concentrations make dissolution harder and reduce the amount of DNA you get back, so if gel extraction is the goal, keep your gel at 1% or below when possible.
Agarose vs. Polyacrylamide
Agarose gels work well for separating DNA fragments roughly from around 100 bp up to tens of thousands of base pairs, depending on concentration. For very small fragments (under about 100 bp) or when you need single-nucleotide resolution, polyacrylamide gels are the better tool. Polyacrylamide forms a much tighter, more uniform pore network and can resolve fragments differing by just one or two bases, which is why it is the matrix used for sequencing gels and applications like microsatellite genotyping.
However, polyacrylamide brings trade-offs. The unpolymerized acrylamide monomer is a potent neurotoxin and needs to be handled with gloves under a fume hood. Polyacrylamide gels are also harder to pour (they require chemical polymerization with TEMED and ammonium persulfate rather than simple heating and cooling) and are typically cast in vertical formats rather than the horizontal trays used for agarose. For most routine molecular biology work, agarose is the default choice because it is safe, cheap, fast, and good enough for the vast majority of fragment-size ranges researchers encounter.
Agarose gels can also separate molecules beyond DNA. They have been used for protein separations under native conditions and for analyzing polysaccharides. Researchers have even used agarose gel electrophoresis to determine the molecular mass of hyaluronan, a large polysaccharide, across a wide size range, though very high-molecular-mass species compress and run together at the top of the gel, just as oversized DNA fragments do.10PubMed Central. Agarose and Polyacrylamide Gel Electrophoresis Methods for Molecular Mass Analysis of 5–500 kDa Hyaluronan
Separating Very Large DNA with Pulsed-Field Gels
Standard agarose gel electrophoresis hits a ceiling for DNA above roughly 20,000–50,000 base pairs. Past that size, DNA molecules are so large that they thread through the gel pores in a stretched-out, snake-like fashion, and all of them migrate at roughly the same speed regardless of size. You lose resolution entirely.
Pulsed-field gel electrophoresis (PFGE) solves this by alternating the direction of the electric field during the run. When the field switches, DNA molecules have to reorient before they can start migrating in the new direction. Smaller molecules reorient faster; larger molecules take longer. In most PFGE systems the reorientation angle is greater than 90°, which forces molecules to briefly move backward before heading in the new direction. Larger molecules retreat farther, so over many switching cycles they fall progressively behind smaller ones.11Cell and Tissue Biology. Pulsed field gel electrophoresis: Theory, instruments and application By tuning the pulse time and electric field strength, researchers can resolve DNA molecules up to several million base pairs, including entire chromosomes from bacteria and yeast.12PubMed. Reorientation time of DNA molecules in pulsed-field gel electrophoresis
PFGE requires specialized equipment (a power supply that can alternate fields and a gel chamber designed for the geometry) and long run times, often 12 to 24 hours or more. The gels themselves are made from standard agarose in TBE, typically at low concentrations (around 1%), but the sample preparation is quite different from routine work. Because the DNA is so large and fragile, cells are embedded directly in agarose plugs, lysed in situ, and the plugs are loaded into the gel wells. Any mechanical shearing during pipetting would break megabase-sized DNA into random fragments and ruin the experiment. PFGE remains the gold standard for bacterial strain typing in outbreak investigations and for constructing physical maps of genomes.
Small Details That Matter More Than You Think
A few easily overlooked points can make a real difference in gel quality. First, always use molecular-biology-grade agarose for electrophoresis. Food-grade or general-lab agarose (the kind used for bacterial plates) contains impurities like sulfated polysaccharides that interfere with DNA migration and enzyme activity. The price difference is modest and the performance gap is large.
Second, the volume of sample you load into each well matters. Overloading a well causes bands to compress, streak, or spill into adjacent lanes. For a standard mini-gel with wells about 5 mm wide, loading 5–10 µL per well with DNA concentrations in the range of 50–200 ng for your band of interest usually gives clean results. If you need to load more volume, use a comb with wider teeth or cast a thicker gel.
Third, pay attention to your DNA ladder. A molecular-weight marker is only useful if it covers the size range of your fragments and if the ladder bands are sharp and well-separated. If your ladder bands look smeary or faint, the ladder may have degraded from too many freeze-thaw cycles or improper storage. Keep ladder aliquots at –20°C and avoid repeated thawing.
Finally, gel imaging conditions make a surprisingly big difference. For ethidium-bromide-stained gels, a short-wavelength UV transilluminator gives the brightest signal but also damages DNA rapidly, which matters if you plan to excise a band. A blue-light transilluminator paired with SYBR-family dyes provides good sensitivity with less DNA damage and is increasingly the preferred setup in labs that do a lot of gel extraction. Whatever system you use, take multiple exposures: an overexposed image hides faint bands, while an underexposed one makes everything look weak.