Poly-L-Lysine Coating for Cell Culture

Poly-L-lysine is a homopolymer of the amino acid L-lysine. At culture pH its side-chain amino groups are protonated, giving the molecule a strong net positive charge. Adsorbed onto glass or onto plasma-treated polystyrene, it converts the surface into a densely cationic layer. The plasma membrane of a mammalian cell carries a net negative charge, contributed by sialylated glycoproteins and glycolipids and by anionic phospholipids, so the cell is held to the surface by simple electrostatic attraction.

That mechanism defines both the usefulness and the limits of the coating.

What it gives you. Attachment where there would otherwise be none or very little, achieved cheaply, reproducibly, with a chemically defined and animal-origin-free material that has no lot-to-lot biological variability. It works on glass, which is otherwise a poor substrate for most cultured cells, which is why it is the default treatment for coverslips destined for imaging.

What it does not give you. Signalling. Integrin engagement by an extracellular matrix ligand triggers focal adhesion assembly, cytoskeletal organisation and downstream survival and differentiation pathways. A charge interaction triggers none of that. Cells on poly-L-lysine alone attach, but many of them spread poorly, and cell types that depend on matrix-derived survival signals — primary neurons, endothelial cells, many stem and progenitor populations — do not thrive on it as a sole substrate.

This is why the common arrangement in neuronal culture is poly-L-lysine or poly-D-lysine first, laminin second: the polylysine layer immobilises the laminin and improves its coverage, and the laminin supplies the receptor-mediated signal.

Poly-lysine is sold in a range of molecular weight fractions, typically from about 30,000 to over 300,000 Da, as a lyophilised powder (usually the hydrobromide salt) or as a ready-made sterile solution.

The trade-off is straightforward. Higher molecular weight chains carry more binding sites per molecule and adsorb more tenaciously, but the solutions are more viscous, harder to pipette evenly and harder to rinse off completely. Lower molecular weight fractions are easier to handle and rinse but bind less firmly.

Published protocols cluster in two places:

• 0.01% (0.1 mg/mL) solutions of the 70,000-150,000 range for routine coating of tissue-culture plastic and coverslips. This is the concentration in the widely used supplier protocol.
• 0.1 to 1 mg/mL of the 30,000-70,000 range in 0.15 M borate buffer, pH 8.3, for glass coverslips in primary neuronal culture, where the alkaline buffer improves adsorption to glass.

Ready-made sterile 0.01% solutions remove the weighing, dissolving and filtering steps and are worth the cost for occasional users. If you make your own, dissolve the powder in sterile tissue-culture-grade water, filter through a 0.22 micron membrane, and store aliquots frozen; the solution is stable frozen but repeated freeze-thaw degrades it. Do not autoclave polylysine solutions.

Work aseptically in a biosafety cabinet throughout. Volumes below follow the standard supplier protocol of 1.0 mL of 0.01% solution per 25 cm2 of growth area, which is approximately 5 micrograms of polylysine per cm2.

1. Prepare the working solution. Dilute stock to 0.01% (0.1 mg/mL) in sterile tissue-culture-grade water. As a reference point, the classical formulation is 5 mg of poly-lysine dissolved in 50 mL of sterile water.
2. Dispense onto the surface. Add 1.0 mL per 25 cm2 — approximately 0.5 mL per well of a 24-well plate, 1 mL per well of a 6-well plate, 3 mL for a T-75 — and rock so the entire growth surface is covered. Uneven coverage at this step is the commonest cause of patchy attachment later.
3. Incubate. Leave at room temperature. 5 minutes is sufficient in the standard protocol; many laboratories use 30 minutes to 1 hour, or overnight at 4 degrees C, without harm. Longer incubation does not increase the adsorbed layer indefinitely, because adsorption saturates.
4. Aspirate the solution completely. Do not let it evaporate in place — evaporation deposits the whole solute load as a crust rather than a monolayer.
5. Rinse thoroughly with sterile tissue-culture-grade water. This step is not optional and is the one most often skimped. Free, unadsorbed polylysine left on the surface is cytotoxic; it permeabilises membranes at concentrations well below those used for coating. Rinse at least three times, and more for high molecular weight material or for sensitive primary cultures.
6. Dry. Aspirate the final rinse and leave the vessel open in the cabinet with the laminar flow running until the surface is completely dry — at least 2 hours. Cells are seeded onto a dry coated surface.
7. Use or store. Coated vessels can be used immediately, or sealed and stored dry. Coated plates keep for weeks to months when stored dry, sterile and protected from light; validate the storage period in your own hands rather than assuming it.

Sterility note: if the polylysine solution was filter-sterilised and the work was done in a cabinet, no further sterilisation is needed. Where UV sterilisation of coated coverslips is used, 10-15 minutes is typical, but prolonged UV exposure damages the coating and the plastic.

Glass needs more preparation than plastic and primary neurons are less forgiving, so this variant is more demanding.

1. Clean the coverslips. Acid-wash (commonly nitric or hydrochloric acid), then rinse exhaustively in distilled water, then in ethanol, and sterilise by baking or autoclaving. Uncleaned coverslips coat unevenly and neurons detach in sheets during medium changes.
2. Prepare polylysine in borate buffer. 0.1 to 1 mg/mL poly-L-lysine (MW 30,000-70,000) in 0.15 M borate buffer, pH 8.3. The alkaline buffer deprotonates surface silanols on the glass and improves adsorption.
3. Cover the coverslips with the solution — around 120 microlitres per 12 mm coverslip is enough if applied as a droplet, or submerge them in a dish.
4. Incubate at least 30 minutes at room temperature, or overnight at 4 degrees C. Overnight is the more common choice for neuronal work.
5. Rinse exhaustively. For primary neurons, rinse ten times or more with sterile distilled water. This is deliberate over-rinsing: residual free polylysine is the leading cause of poor neuronal survival in the first 48 hours after plating, and neurons are far more sensitive to it than cell lines.
6. Dry in the cabinet, and store dry and sterile.
7. Add laminin if the protocol calls for it. Apply laminin at 5-50 micrograms/mL (commonly around 10 micrograms/mL for neurons) in PBS or in the base medium, 2 hours at 37 degrees C or overnight at 4 degrees C. Do not rinse the laminin off and do not let it dry — aspirate immediately before seeding and add the cell suspension straight away. Dried laminin loses activity, and laminin-coated surfaces are best used fresh rather than stored.

The two are enantiomeric polymers of the same amino acid and are chemically and electrostatically equivalent. The difference is biological.

Proteases secreted by cultured cells cleave peptide bonds between L-amino acids and do not cleave bonds between D-amino acids. Poly-L-lysine is therefore slowly degraded by the cells growing on it; poly-D-lysine is not. Two consequences follow:

• Poly-D-lysine is preferred for long-term culture, particularly primary neurons and glia maintained for two weeks or more, where a degrading substrate progressively loses adhesion and releases free lysine oligomers into the medium.
• Poly-L-lysine is entirely adequate for short-term work — attaching a cell line for a 48-hour experiment, immobilising cells for imaging, or cytospin preparations — and it costs less.

The commonly repeated concern that poly-L-lysine degradation releases toxic quantities of free L-lysine is worth stating carefully: the degradation is real, but the practical problem in most failed cultures is not lysine toxicity, it is loss of adhesion over time combined with inadequate rinsing of free polymer at the coating step. If neurons are dying in the first two days, suspect rinsing. If a two-week culture lifts off in week two, suspect the L isomer.

Where the culture is long, the cells are primary, or the experiment is expensive, use poly-D-lysine. Otherwise poly-L-lysine is a reasonable default.

Most established adherent cell lines — HeLa, CHO-K1, Vero, most fibroblast lines — attach perfectly well to standard tissue-culture-treated polystyrene, which is already plasma-treated to bear negatively charged carboxyl and hydroxyl groups and which adsorbs attachment proteins from the serum in the medium. Coating them is unnecessary work and an unnecessary variable.

Coating is genuinely needed when:

• The substrate is untreated glass or untreated plastic. Coverslips for microscopy, chamber slides, some optical-bottom plates, and microfluidic devices.
• The cells are weakly adherent. HEK293 is the standard example: it grows on plastic but lifts in sheets during medium changes and washes, and a polylysine coating largely solves this.
• The culture is serum-free. Without serum there are no adsorbed attachment proteins, so the surface has to supply the adhesion itself.
• The cells are primary, neural, or stem or progenitor populations that require a defined adhesive substrate.
• The protocol involves repeated washing — immunocytochemistry, in-cell assays, imaging with multiple medium exchanges — where marginal adhesion becomes cell loss.
• You need a defined, animal-origin-free surface for regulatory or reproducibility reasons, where polylysine's synthetic nature is an advantage over animal-derived matrix proteins.

Coating is the wrong answer when the real problem is something else. Cells that will not attach because they were over-trypsinised, because viability is low, because the medium is wrong or because the culture has mycoplasma will not attach to a coated surface either. Check viability and passage history before reaching for a coating.

When charge alone is not enough, the alternatives are real extracellular matrix proteins that engage integrin receptors.

Laminin is a large heterotrimeric glycoprotein of the basement membrane and the principal substrate for neurons, neural progenitors and many endothelial cells. Published protocols use 5-50 micrograms/mL, commonly around 10 micrograms/mL for neuronal work, in PBS or base medium, applied for 2 hours at 37 degrees C or overnight at 4 degrees C. Laminin coatings should be used fresh and must not be allowed to dry; the solution is aspirated immediately before seeding and is generally not rinsed off. Laminin is heat-sensitive and should be thawed slowly at 4 degrees C and not refrozen.

Fibronectin is a dimeric plasma and matrix glycoprotein carrying the RGD motif recognised by several integrins. It is the usual choice for fibroblasts, endothelial cells and mesenchymal stromal cells. Typical coating is around 1 microgram/cm2, applied for 1 hour at room temperature or overnight at 4 degrees C, followed by three rinses with sterile water or PBS. Ready-made fibronectin/gelatin blends are applied at 0.2-0.25 mL/cm2 for 2-3 hours at room temperature, then aspirated and washed three times with sterile PBS.

Collagen type I is the standard substrate for hepatocytes, many epithelial cells and smooth muscle. It is supplied acid-solubilised and behaves as a thermosetting gel: collagen types I and IV remain soluble at 4 degrees C, begin to solidify at room temperature and gel quickly at 37 degrees C. That property is exploited to make thin films (thin layer, air-dried) or thick gels (allowed to set at 37 degrees C), which are biologically different substrates. Neutralise and keep the solution cold while dispensing if you want an even thin coating.

Gelatin is denatured collagen: cheap, easy, sterilisable by autoclaving, and lacking the intact triple-helical structure that gives native collagen its integrin-binding properties. It is the standard low-cost coating for embryonic stem cells on feeders and for retroviral packaging lines. Typical surface coverage is around 100-200 micrograms/cm2, which is what a 0.1-0.2% solution delivers at normal working volumes.

For all of these, suppliers state explicitly that the coating concentration should be optimised for the specific cell type. Titrate over at least a three-point range on your own cells before fixing a protocol.

Cells attach, then die within 24-48 hours. Residual free polylysine. This is the dominant failure mode in primary culture and it is entirely a rinsing problem. Increase the number of water rinses substantially — ten or more for neurons — and confirm the polylysine solution was not applied at higher than 0.01% for plastic.

Patchy attachment, cells growing in islands with bare regions between. Uneven coating. Either the solution did not cover the whole surface, the vessel was not rocked, or the solution partially evaporated during a long incubation in a poorly humidified cabinet. Use enough volume to cover completely and rock deliberately.

A visible film, crust or crystalline deposit on the surface. The coating solution was allowed to dry in place instead of being aspirated and rinsed. Discard the vessel; a dried deposit cannot be rescued.

Cells round up, clump or refuse to spread on a freshly coated surface. The coating is too concentrated. Excess cationic charge holds cells so tightly that they cannot remodel their adhesions and spread. Dilute the coating solution and rinse more.

Coating works on plastic but not on glass. Glass needs cleaning and benefits from the borate buffer variant. Untreated, uncleaned coverslips coat unevenly regardless of concentration.

Cells detach after one to two weeks in long-term culture. Proteolytic degradation of poly-L-lysine. Switch to poly-D-lysine.

Laminin coating gives no benefit. Usually because it was allowed to dry, was rinsed off, was stored too long, or was thawed too fast. Coat fresh, keep wet, aspirate immediately before seeding.

Coated plates that worked last month do not work now. Coated surfaces are not indefinitely stable, and polylysine stock degrades with repeated freeze-thaw. Aliquot the stock, date the coated plates, and set a validated shelf life rather than an assumed one.

No coating helps at all. The problem is not the surface. Check viability by trypan blue, check the passage number, check for mycoplasma, and check that the cells were not over-trypsinised at the last passage.