Custom Peptide

Q: How should I dissolve peptides?

A: The solubility of a peptide is determined mainly by its polarity. Acidic peptides can be reconstituted in basic buffers, whereas basic peptides can be dissolved in acidic solutions. Hydrophobic peptides and neutral peptides that contain large numbers of hydrophobic or polar uncharged amino acids should be dissolved in small amounts of organic solvent such as DMSO, DMF, acetic acid, acetonitrile, methanol, propanol, or isopropanol, and then diluted using water. DMSO should not be used with peptides that methionine or free cysteine because it might oxidize the side-chain.

Test a portion of the synthesized peptide before dissolving the rest of the sample. Lyophilized peptides should be centrifuged briefly to pellet all the material. You might need to test several different solvents until you find the appropriate one. Sonication can be used to enhance solubility.

  1. First, assign a value of -1 to each acidic residue (Asp [D], Glu [E], and the C-terminal -COOH). Next, assign a value of +1 to each basic residue (Arg [R], Lys [K], His [H], and the N-terminal -NH2), and then calculate the overall charge of the peptide.
  2. If the overall charge of the peptide is positive, the peptide is basic. Try to dissolve the peptide in distilled water if possible. If it fails to dissolve in water, then try to dissolve the peptide in a small amount of 10-25% acetic acid. If this fails, add TFA (10-50 ul) to solubilize the peptide, and then dilute it to your desired concentration.
  3. If the overall charge of the peptide is negative, the peptide is acidic. Acidic peptides might be soluble in PBS (pH 7.4). If this fails, add a small amount of basic solvent such as 0.1 M ammonium bicarbonate to dissolve the peptide, and then add water to the desired concentration. Peptides that contain free cysteines should be dissolved in de-gassed acidic buffers because thiol moieties will be oxidized rapidly to disulfides at pH >7.
  4. If the overall charge of the peptide is 0, the peptide is neutral. Neutral peptides usually dissolve in organic solvents. First, try to add a small amount of acetonitrile, methanol, or isopropanol. For very hydrophobic peptides, try to dissolve the peptide in a small amount of DMSO, and then dilute the solution with water to the desired concentration. For Cys-containing peptides, use DMF instead of DMSO. For peptides that tend to aggregate, add 6 M guanidine, HCl, or 8 M urea, and then proceed with the necessary dilutions.

To prevent or minimize degradation, store the peptide in lyophilized form at -20 Celsius degree, or preferably -80 Celsius degree. If the peptide is in solution, freeze-thaw cycles should be avoided by freezing individual aliquots.

Positively charged residues: K, R, H, and the N-terminus
Negatively charged residues: D, E, and the C-terminus
Hydrophobic uncharged residues: F, I, L, M, V, W, and Y
Uncharged residues: G, A, S, T, C, N, Q, P, acetyl, and amide

RKDEFILGASRHD: (+5) + (-4) = +1 This is a basic peptide. See step #2 above.
EKDEFILGASEHR: (+4) + (-5) = -1 This is an acidic peptide. See step #3 above.
AKDEFILGASEHR: (+4) + (-4) = 0 This is a neutral peptide. See step #4 above.

Q: How do I choose the best level of peptide purity for my research?
A: Crude peptides are not recommended for biological assays. Crude peptides may contain large amounts of non-peptide impurities such as residual solvents, scavengers from cleavage, TFA and other truncated peptides. TFA cannot be totally removed. Peptides are usually delivered as TFA salt. If residual TFA is a problem for your experiment, we recommend other salt forms such as acetate and hydrochloride. These salt forms are usually 20-30% more expensive than the regular TFA salt. This is due to the peptide loss that takes place during the salt conversion and the greater amounts of raw materials required.
Purity Assays/studies
>70% purity
  • Peptide arrays
  • Antigens for antibody production
  • Competitive elution chromatography
  • ELISA standards for measuring antisera titers
>80% purity
  • Western blotting studies (non-quantitative)
  • Enzyme-substrate studies (non-quantitative)
  • Peptide blocking studies (non-quantitative)
  • Affinity purification
  • Phosphorylation assays
  • Protein electrophoresis applications and immunocytochemistry
>95% purity
  • ELISA standards and RIA protocols (quantitative)
  • Receptor-ligand interaction studies (quantitative)
  • In vitro bioassays and in vivo studies
  • Enzyme studies and blocking assays (quantitative)
  • NMR studies
  • Mass spectrometry
  • Other quantitative assays
>98% purity
  • SAR Studies
  • Clinical trials
  • APIs (Active Pharmaceutical Ingredients)
  • Commercial products
  • X-ray crystallography studies
  • Other sensitive experiments: enzyme-substrate studies, receptor-ligand interaction studies, blocking and competition assays
Q: How should I store and handle my synthesized peptides?

A: Peptides are shipped at room temperature, and are highly stable at lyophilized form in sealed bags. Peptides should not be kept in solution for long periods of time.

Peptide storage guidelines: For long-term storage, peptides should be stored in lyophilized form at -20 Celsius degree, or preferably at -80 Celsius degree with desiccant in sealed containers to minimize peptide degradation. Under these conditions, peptides can be stored for up to several years. This type of storage prevents bacterial degradation, oxidation, and the formation of secondary structures.

Opening the package: It is better to equilibrate the peptides to room temperature in a desiccator prior to opening and weighing. Failure to warm the peptides beforehand can cause condensation to form (peptides tend to be hygroscopic) on the product when the bottle is opened. This will reduce the stability of the peptides.

Before reconstitution, centrifuge the vial of lyophilized peptide at 12,000 x g for 20 seconds. This will help pellet the entire peptide sample for reconstitution.

Weighing peptides: Weigh out your required quantity of peptide rapidly and store all unused peptide at -20 Celsius degree or below. Sequences that contain cysteine, methionine, tryptophan, asparagine, glutamine, and N-terminal glutamic acid will have a shorter shelf life than other peptides.

Q: Can I predict whether a peptide will be soluble?

A: The solubility of a peptide in water cannot be predicted by studying its structure. However, the ε-amino group of Lys and the guanidine of Arg are usually helpful for estimating the solubility of peptides, particularly those with short sequences. In contrast, acidic peptides that contain Asp and Glu tend to be insoluble in water, but can be dissolved easily in diluted ammonia or basic buffers.

Certain basic characteristics can be used to predict solubility:

  • Peptides containing <5 amino acids are commonly soluble in aqueous solutions. However, if the entire sequence consists of hydrophobic residues it will have only limited solubility or could be completely insoluble.
  • Hydrophilic peptides that contain >25% charged amino acids (E, D, K, R, and H) and <25% hydrophobic residues are usually soluble in aqueous solutions.
  • Hydrophobic peptides whose sequence contains at least 50% hydrophobic residues might be completely or only partially soluble in aqueous solutions. These peptides should instead be dissolved in organic solvents such as DMSO if they do not contain C, W, or M residues. If they do contain these amino acids, they should be dissolved in DMF, acetonitrile, isopropyl alcohol, ethanol, acetic acid, 4-8 M guanidine hydrochloride (GdnHCl), or urea prior to being diluted carefully in aqueous solution.
  • Hydrophobic peptides that include >75% hydrophobic amino acids are generally not soluble in aqueous solutions. Instead, very strong solvents such as TFA or formic acid must be used for the initial solubilization. However, the peptide might precipitate when added to an aqueous buffered solution. As such, high concentrations of organic solvents or denaturants might be needed to dissolve these peptides.
  • Peptides that include a very high proportion (>75%) of D, E, H, K, N, Q, R, S, T, or Y can form intermolecular hydrogen bonds (cross-links), which can result in gel formation in concentrated aqueous solutions. Therefore, peptides should be dissolved in an organic solvent that is compatible with the final experiment. After dissolving the peptides in organic solvent, the solution should be added slowly (dropwise) to a stirring aqueous buffered solution. The limit of solubility is reached when the resulting peptide solution begins to show turbidity.
Q: What salt form should I use?
A: Peptides are usually delivered as TFA salts. If residual TFA would be problematic for your experiment, we recommend other salt forms such as acetate and hydrochloride. These salt forms are usually 20-30% more expensive than the regular TFA salt because of the peptide loss that takes place during the salt conversion and the greater amounts of raw materials required.
Q: What are acetylation and amidation?

A: Chemically synthesized peptides carry free amino and carboxy termini. The need for N-terminal acetylation or C-terminal amidation must be stated explicitly during ordering. It is impossible to perform these modifications after synthesis has been completed.

N-terminal acetylation and C-terminal amidation reduce the overall charge of a peptide and decrease solubility. However the stability of the peptide usually increases because the terminal acetylation and amidation allow the peptide to mimic the native protein more closely. In this way, these modifications may increase the peptide's biological activity.

Q: Is a spacer required for fluorescent modification?

A: Usually, dyes such as biotin and FITC can be introduced either N-terminally or C-terminally. We recommend N-terminus modification for its higher success rate, shorter turnaround time, and ease of operation. Peptides are synthesized from the C-terminus to the N-terminus. N-terminus modification is the last step in the SPPS protocol. No more specific coupling steps are required. In contrast, the C-terminus modification requires additional steps and is usually more complex.

Most dyes are large aromatic molecules. The incorporation of such bulky molecules may help to avoid interactions between the label and the peptide. This will help maintain peptide conformation and biological activity. It is recommended that a flexible spacer such as Ahx (a 6 carbon linker) be included to render the fluorescent label more stable. Otherwise, FITC could easily link to a cysteine thiol moiety or the amino group of lysine at any position.

Q: How do I Detect Small Peptides using SDS-PAGE?

A: If your sample contain proteins of interest that are <20 kDa, please download a protocol that explains how to detect synthetic peptides using SDS-PAGE, including effective methods for Coomassie blue staining, silver staining, and electroblotting.

Tricine-based SDS-PAGE is used most commonly to separate proteins sized 1-100 kDa, and is the electrophoretic system of choice for resolving proteins <30 kDa. Although visualizing small peptides using SDS-PAGE is challenging, Tris-tricine gels afford better resolution. However, if you simply want to detect the peptide, MS remains the most accurate method for confirming the identity of a peptide.

Small peptides binds to Coomassie brilliant blue less readily that do larger proteins. Therefore, smaller peptides are difficult to detect using Coomassie or silver staining. Additional sample could be loaded to allow peptides to be visualized on gels; changing the percentage of the gel will only help if you think that your peptide migrated out gel. In this instance, the percentage of cross linker in a regular 17% gel could be increased, and the pH of the resolving gel could be increased to 9.5 (compared with the normal 8.8). Finally, the addition of 4-8 M urea helps sharpen bands.

The use western blotting rather than gel staining is a far more sensitive detection method. However the peptide might simply pass through the membrane during transfer. If you think this occurs, the experiment can be repeated using two pieces of membrane and shorter transfer time (<1 hour at 200 mA). A membrane with a 0.2-um pore size should be sufficient: although smaller pore sizes are available, they should not be necessary. An additional option would be to try semi-dry transfer for 15-20 minutes using the current density (mA/cm2) recommended for the apparatus. A short transfer time of 15 min works for most small peptides. If it is possible to plan ahead, a control small peptide labeled with biotin could be synthesized to monitor the transfer process and assess the ability of the peptide to bind to the membrane using streptavidin-conjugated HRP.

Q: How do I dissolve peptides in DMSO?

A: Dimethyl sulfoxide (DMSO) is an organosulfur compound with the formula (CH3)2SO. DMSO is used frequently in cell banking applications as a cryoprotectant because it prevents intracellular and extracellular crystals from forming in cells during the freezing process. For most cryopreservation applications, DMSO is used at a concentration of 10%, and is usually combined with saline or serum albumin.

Hydrophobic peptides can be dissolved easily in DMSO. However, peptides in DMSO might be cytotoxic to cells, even though DMSO increases cell permeability. High concentrations of DMSO should never be used for cell culture. 5% is very high, and will dissolve the cell membranes. Most cell lines can tolerate 0.5% DMSO, and some cells can tolerate up to 1% without severe cytotoxicity. However, primary cell cultures are far more sensitive. Therefore, if you are using primary cells a dose-response curve (viability) should be performed using DMSO concentrations <0.1%.

Try to dissolve very hydrophobic peptides in a small amount of DMSO (30-50 ul, 100%), and then slowly add the solution drop-wise to a stirring aqueous buffered solution such as PBS (or your desired buffer) to the required concentration. If the resulting peptide solution begin to show turbidity, you have reached the limit of solubility. Sonication will help dissolve the peptides.

Rule of thumb:

  • 0.1% DMSO is considered to be safe for almost all cells.
  • A final concentration of 0.5% DMSO is used widely for cell culture without cytotoxicity.
  • 1% DMSO does not cause any toxicity in some cells, but 0.5% DMSO is recommended.
  • 5% DMSO was used successfully in some cells.
  • To maintain a final concentration of 0.5%, you can make 200x stock in 100% DMSO.
Q: What are your QC standards for peptide synthesis?
A: All materials supplied to NovoPro are considered the confidential property of the customer. NovoPro provides free HPLC and MS results with your package. Peptides are purified by reverse-phase chromatography. The chromatogram indicates the number and relative amount of by-products. The molecular mass of the peptide is determined by mass spectrometry to confirm that the correct product is being delivered. MS results also show the masses of the main impurities. Additional analysis revealing net peptide content can be performed upon request. Net peptide content is indicated by either amino acid analysis or elemental analysis. These methods allow the verification of the amino acid composition of the peptides. They serve as additional means of confirmation of peptide identity. All synthetic peptides meeting the customer's purity criteria are sent. All residual materials, such as peptides not meeting the customer's purity criteria are discarded. These residual materials can be sent to the customer upon request.
Q: What is net peptide content?
A: The net peptide content is different from the peptide purity. The net peptide content is the percentage of peptides relative to nonpeptidic materials, mostly counterions and moisture. The net peptide content can be determined by amino acid analysis. Please place a request for a quote if you require this service. Usually, hydrophilic peptides absorb tiny amounts of moisture even under strict lyophilization conditions. Net peptide content may vary from batch to batch depending on the purification and lyophilization processes.