Laboratory solution preparation
Planning a multi-component peptide solution
This page helps you work out, in the lab or the classroom, how much of each ingredient to pour when you want one solution that holds several peptides, such as a calibration mix for an instrument, a pool for a cell assay or an exercise for a teaching class. You tell the mixer what you have and what you want, and it does the arithmetic and writes the answer in plain sentences. Below the mixer you will find a plain-language explainer and common questions, and further down a fuller laboratory planner for trained researchers.
For laboratory research use only. Not for human or veterinary use.
On this page
Solution mixer
This tool calculates concentrations for laboratory solutions. GPC gives no administration advice.
General laboratory information.
There is very little research on how different peptides behave when mixed into one solution. Mixing can speed up breakdown or change how they behave. Treat this as arithmetic, and measure what you make.
In the picture
Example: three ingredients at 5, 5 and 10 mg per mL, each wanted at 0.1 mg per mL in 1 mL. Pour in 20.0 µL, 20.0 µL and 10.0 µL, then top up with 950.0 µL of liquid.
How the mixer thinks
Every ingredient starts in its own liquid, at its own strength. You pour a measured amount of each into one container, then top up with plain liquid to the final amount you want.
Strength just means how much of the ingredient is in each mL of its liquid, and the mixer writes it in mg per mL. Its job is to work out how much of each liquid to pour so that every ingredient ends up at the strength you asked for.
A picture with ink:
- Imagine a bottle of strong blue ink. Pour a little into a jar, then fill the jar with clear water up to the line. The jar is now pale blue, but you did not add any more blue. You only spread the same blue through more liquid. Top-up liquid always works this way: it never adds more of an ingredient, it only spreads it out.
The rule behind it is short. The strength of an ingredient in the final mix is how much of it you poured in, divided by the final amount of liquid. Pour the same amount into a bigger final amount and the strength goes down. Pour more into the same final amount and it goes up.
So the mixer runs that sum backwards. You say what strength you want for each ingredient and what final amount of liquid you want, and it works out how much of each solution to pour. Whatever room is left in the container is filled with plain top-up liquid.
You choose one of three ways to start, in the first step: start with an empty container, combine what you have already dissolved, or add to a container that already has liquid in it.
- The rule
- Strength in the final mix = how much of the ingredient you poured in ÷ the final amount of liquid
Why some plans do not fit
If your ingredients are too weak to reach your targets, the pours add up to more than the container can hold.
Think of paint. If your paint is pale, you need a great deal of it to make a deep colour, so much that the tin has no room left for anything else. Here the container is the tin, and each ingredient needs its own share of the room.
A tiny worked example with made-up ingredients:
- A plan that fits. You want Ingredient A, Ingredient B and Ingredient C each at 0.1 mg per mL in a final amount of 1 mL. They start at 5, 5 and 10 mg per mL. A needs 20 µL, B needs 20 µL and C needs 10 µL. That is 50 µL in all, so 950 µL of top-up liquid fills the rest. There is plenty of room to spare.
- A plan that does not fit. Now you want each at 1 mg per mL in 1 mL, starting from weaker solutions at 2, 2 and 5 mg per mL. A needs 0.5 mL, B needs 0.5 mL and C needs 0.2 mL. That is 1.2 mL of pours for a container that holds 1 mL. There is no room.
Making the final amount bigger does not rescue the second plan. At 2 mL, the same targets need 1, 1 and 0.4 mL, which is 2.4 mL, still 20% too much. Every pour grows along with the container, so the plan stays 20% over at any size.
Two things do help. One is a stronger solution: if A and B can be dissolved at 4 mg per mL, the pours become 0.25, 0.25 and 0.2 mL, which is 0.7 mL, and 0.3 mL is left for top-up liquid. The other is a lower target: asking for 0.5 mg per mL of A and B gives the same pours. Leaving an ingredient out also frees up room.
- Quick check
- For each ingredient, divide its target by its strength. Add those shares up. If the total is more than 1, the plan does not fit.
Things worth knowing before you mix
Every ingredient arrives in its own liquid, and a mix inherits all of them. The mixer does the arithmetic. It cannot tell you what will happen inside the liquid, so these are the things to keep in mind.
Acids and bases can clash
Some peptides dissolve best in a little acid, and others in a little base. Bachem dissolves basic peptides first in acetic or trifluoroacetic acid, and acidic peptides first in a small amount of a basic liquid such as 0.1% ammonia in water. If both end up in one container, the acid and the base cancel each other out (chemists say they neutralise), and the final acidity lands somewhere between the two. Measure it rather than guess. There is one more trap. Pouring an acid onto a liquid made with ammonium bicarbonate releases carbon dioxide gas, and bubbles and foam can bend proteins out of shape (Hedges et al., 2013). It is better to pick liquids that do not pair an acid with a bicarbonate.
Different peptides like different acidity
Acidity is measured on the pH scale. Every peptide dissolves least well at one particular pH, called its isoelectric point (the pH at which it carries no net charge). For peptides that are often combined, the estimated points run from about 3.9 to 9.7, so no single pH suits every ingredient. Such estimates can be off by more than one pH unit (Shaw et al., 2001), which is one more reason to measure.
Some liquids add up across ingredients
Some ingredients are dissolved in DMSO (dimethyl sulfoxide, a common laboratory solvent). Each one brings its share of DMSO into the final mix, so the total grows with every ingredient that carries it. Two ingredients in 16% DMSO, each poured at 1 part in 100, bring 0.16% each, or 0.32% together. Cell experiments usually keep the total at or under 1% by volume, and JPT and STEMCELL Technologies both give that limit. A bigger final amount does not lower the total if every strength stays the same, because every pour grows with it.
Some ingredients are fragile
Some ingredients react easily with oxygen, which changes them. This is most true of ones with sulfur-containing parts (the amino acids cysteine and methionine) and of tryptophan. JPT advises against DMSO for peptides containing methionine or cysteine, because it can change the side chain. The reaction is faster at high pH and during freeze–thaw cycles (Hoofnagle et al., 2016), and Bachem notes that free cysteine reacts rapidly above pH 7. Copper-bound ingredients are another case, because copper can pass to other molecules and can react with sulfur-containing ones in a test tube (Eben and Imlay, 2023). We found no study showing this between peptides in a mixture, but it is best to keep these ingredients apart until they have been tested.
Thin solutions lose more to the container walls
The more you thin a solution, the larger the share that can stick to the wall of the container, and it does not stick evenly across ingredients. In one supplier’s calibration mix, three of the fifteen peptides gave a signal that was 20 to 60% lower, and more variable, when kept at the lower strength. If you have to thin a mix, do it close to the time your method runs.
Very little is known about peptides mixed together
There has been very little formal research on whether different peptides stay compatible and stable when they are mixed into one solution. Mixing can speed up the breakdown of one of them or change how it behaves, through a shift in acidity, a shared liquid, or the peptides reacting with each other. We looked too. A literature search on 27 September 2026 found no study describing the solution chemistry of a combination. That does not show a mix is stable. It means the question is open, which is why labs measure a mixture, for example by liquid chromatography (HPLC) or mass spectrometry, instead of assuming.
These are cautions taken from supplier guidance and method papers. A mix that raises none of them has not been shown to be compatible.
What the colours and the vial show
Each ingredient gets its own colour, and the vial fills in the order you pour, with the pale top-up liquid last.
Think of layers of dye in a clear jar. The first ingredient sits at the bottom in its colour, the next one goes on top in its colour, and so on. Each coloured layer is the share of the container that one pour takes up. The pale layer at the top is the plain liquid that fills whatever room is left.
The picture is a guide, not a measurement. Measure your real pours with your own calibrated tools, and read the numbers the mixer writes in sentences, not the height of a layer.
Common questions about the mixer.
Can I mix more than four things?
Both planners on this page handle up to four ingredients at a time. For a bigger mixture, plan it in stages: prepare one mixture of up to four, then treat it as a single ingredient in the next plan. The laboratory planner further down also lets you enter molecular weights and the finer details a trained researcher would want.
Why does it say my plan does not fit?
Because your ingredients are too weak to reach your targets inside the container you chose. Each ingredient takes up a share of the room, and together the shares come to more than the whole container. Making the final amount bigger does not fix this, because every pour grows with it. A stronger solution, a lower target or one less ingredient does.
Do I need molecular weights?
Not for the basic mixer. Strengths in mg per mL are enough. Molecular weights only matter when you want an equal number of molecules of each peptide, and the laboratory planner below handles that.
Is it okay to mix peptides together?
It depends, and honestly nobody can say for sure yet. There has been very little research on how different peptides behave when they share a solution. Mixing can speed up the breakdown of one of them or change how it behaves, through acidity, a shared liquid or the peptides reacting with each other. This tool only does arithmetic. It cannot tell you whether a mix is stable, so test your mixture, for example by liquid chromatography or mass spectrometry, before you rely on it.
Why is the top-up amount so large?
Usually because your ingredients are strong. A small pour of a strong solution spreads out into a much larger final amount, and the top-up liquid fills the rest. In the example above, 50 µL of pours leave 950 µL of top-up liquid in a 1 mL container. A large top-up is a good sign: it means there is plenty of room.
What if my powder is not dissolved yet?
The basic mixer can start there. Enter the amount of powder you have and the amount of liquid you dissolved it in, and it works out the strength of that solution for you.
Need more detail? The advanced laboratory planner
The planner below is for trained researchers who want full control. It does everything the mixer above does, and then adds the detail a method needs.
Here is what it adds:
- Molecular weights for each component, so it can show strengths in µM beside mg per mL, which is what you need for an equimolar mix (the same number of molecules of each peptide).
- Amounts entered by mass or by moles, whichever your method uses.
- Solvent flags. You mark which stock solutions are in DMSO and at what percentage, and it adds up the total from every one of them.
- Three preparation modes with their laboratory names: stocks into an empty container, combining whole preparations, and adding to an existing solution.
- Capacity checks on every transfer and on the container, such as a warning for a transfer that is too small to pipette well or a container filled above 90% of its capacity.
- A flags list that points out the chemistry worth checking, from acid meeting bicarbonate to DMSO with oxidation-prone ingredients and copper-bound ones.
The flags work by rule, to show you where to look. They do not predict what will happen in the solution. The explanations after the planner go into the laboratory detail.
Multi-component solution planner
Advanced laboratory planner
This tool calculates concentrations for laboratory solutions. GPC gives no administration advice.
General laboratory information.
We found no published study of compatibility or stability for common combinations in solution. Mixing can degrade a component or change its behaviour. The component with the shortest stability in solution governs the mixture. A fixed mixture removes single-component controls, so an observed response cannot be assigned to one component. Verify a prepared mixture by HPLC or MS when the distinction matters. This planner provides arithmetic for laboratory education only.
An informal peptide nickname can refer to a short acetylated fragment or a full-length chain. The identity stated on a certificate of analysis governs; the nickname alone does not identify the material.
Illustration key
Where are multi-peptide solutions used in the laboratory?
In two well-established places: peptide pools for cell-based immunoassays, and mixed calibration standards for LC-MS. In both, the amount is stated for each peptide separately.
When a mixture is documented, you get a concentration or an amount for each peptide, never one figure for the whole. The planner works the same way: it reports every component separately.
Here are two examples from supplier documents:
- Peptide pools. JPT supplies its standard pools with 25 µg, or 15 nmol, of each peptide. STEMCELL Technologies makes up such a pool as a stock at 100 µg/mL per peptide in 16% (v/v) DMSO (dimethyl sulfoxide, a common laboratory solvent). It generally recommends a final concentration of at least 1 µg/mL per peptide in the assay, with DMSO kept below 1% (v/v).
- Calibration mixtures. Thermo Scientific’s peptide retention time calibration mixture holds 15 isotope-labelled peptides, each at 0.5 or 5 pmol/µL. Its user guide describes the mixture as a tool for checking how well an instrument performs, and uses at least 8 to 10 of the peptides to calibrate the column.
Method papers build larger sets in the same way. Hoofnagle and colleagues, writing about peptides for mass spectrometry-based assays, prepared two mixtures of 50 peptides, at 200 and at 1000 fmol/µL of each peptide.
An equal-mass pool and an equimolar pool are two different mixtures, so it helps to decide which one you want first. Equal mass means the same µg/mL of every peptide, so the number of molecules differs with each peptide’s molecular weight. Equimolar means the same µM of every peptide, as in the calibration mixture above, and it needs each component’s molecular weight and its net amount. When you enter a molecular weight for a component, the planner shows µM beside mg/mL for it.
In a cell-based assay, the DMSO that each stock carries in quietly adds up. A 1:100 dilution of a stock in 16% DMSO adds 0.16% (v/v) to the final solution, and two such stocks add 0.32%. The planner totals the co-solvent from every stock you mark as DMSO, at the percentage you enter, so you do not have to keep count yourself.
- 25 µg (15 nmol)
- Of each peptide in a standard JPT peptide pool
Why does a fixed mixture limit what an experiment can show?
Without single-component controls, you cannot tell which component is behind a measured response.
Statisticians have a name for this. When one estimate carries the influence of several factors at once, the factors are called aliased: the data cannot pull them apart. The NIST/SEMATECH handbook of statistical methods defines the term for designed experiments. A fixed mixture tested on its own is the extreme case, because every component changes together and each one is aliased with all the others.
A picture with ink:
- Mix blue ink and yellow ink into green, and let a drop of the green stain a test strip. The strip cannot tell you whether the blue or the yellow made the stain. To find out, you need one strip with blue alone and one with yellow alone. A fixed mixture is the green ink without the single-colour strips.
A design that can tell the components apart keeps a separate arm (its own set of samples) for each one, at the concentration that component has in the mixture. Preparing the mixture from separate stocks makes those arms possible. The planner reports each component’s final concentration, which is exactly what its own arm needs.
The ratio becomes a variable too. A mixture at 1:1:1 by mass is not the same as one at 1:1:1 by moles, and a reading from one tells you nothing about another ratio. Write down the ratio and what it is based on, and keep that note with the solution.
A single purity figure is not defined for a mixture either. Bachem defines HPLC purity as the percentage of the target peptide compared with impurities that absorb light at 220 nm, so the figure belongs to one target peak. For a mixture, you want an identity check and a peak-area reading for each component.
What is known about peptides combined in solution?
There has been very little formal research on whether different peptides stay compatible and stable when they are mixed into one solution. Mixing certain peptides can quickly break one of them down or change how it behaves. Some ready-made peptide blends are popular, but putting different peptides into a single solution still carries real risks: a change in pH, or different amino-acid chains reacting with one another in ways that cause degradation or change how they behave.
We looked for answers too. A literature search on 27 September 2026 paired the peptides most often combined with one another with the words stability, degradation, mixture and compatibility. It returned reviews and studies about other questions, and none described the solution chemistry of a combination. It is worth saying plainly: a lack of published data does not show that a combination is stable. It means the question is open.
What we do know is general, and it points the same way. Bachem advises against storing peptide solutions for long, especially when the peptide contains Asn, Gln, Cys, Met or Trp (shorthand for the amino acids asparagine, glutamine, cysteine, methionine and tryptophan). In a mixture, the component that keeps least well in solution sets the limit for the whole solution.
Mixing can also break a component down or change how it behaves. That can happen through the pH the mixture settles at, through a co-solvent that another stock brings in, or through reactions between the components themselves. The next section lists the combinations to check.
If the composition of a prepared mixture matters to your work, measure it rather than assume it. HPLC or MS (mass spectrometry) confirms that each component is present and intact. Amino acid analysis is the reference method for concentration, benchmarked with UV absorbance (Hoofnagle et al., 2016). Our teaching guide covers running peptides by HPLC. And if what you want to know is what a sealed pre-blended powder contains, our testing guide explains how to send the original sealed vial to an outside laboratory, which cannot accept a solution you prepared.
- Asn, Gln, Cys, Met, Trp
- Residues that make long-term storage of a peptide solution especially inadvisable (Bachem)
What chemistry should be checked before stocks are combined?
Every stock arrives with its own solvent and its own pH, and the mixture inherits all of them. This is the tricky part, and it pays to take it slowly. The planner flags these combinations by rule so you know where to look; it does not predict what will happen in the solution.
One pH has to serve every component
Bachem dissolves basic peptides first in acetic or trifluoroacetic acid, and acidic peptides first in a small amount of a basic solvent such as 0.1% aqueous ammonia. So the stocks of one mixture can sit at opposite ends of the pH scale: 10% (v/v) acetic acid is near pH 2.3, and 0.1% is near pH 3.3. When they are combined, acid and base neutralize each other, and the final pH lands between them, at a value set by the amounts and by how strongly each stock resists a change in pH (its buffer capacity). Measure it rather than assume it. A peptide dissolves least well near its isoelectric point, the pH at which it carries no net charge, and the estimated isoelectric points of commonly combined peptides span about 3.9 to 9.7, so no single pH suits every component. Such estimates can be off by more than one pH unit (Shaw et al., 2001).
Carbon dioxide and foam
Adding an acid stock to a stock dissolved in ammonium bicarbonate releases carbon dioxide. Bubbles and foam create gas–liquid interfaces, and proteins that stick to such an interface go through major changes in shape, or conformation (Hedges et al., 2013). Where the sequences allow it, choosing stocks that do not pair an acid with a bicarbonate avoids the problem.
DMSO adds up across stocks
Each DMSO stock carries its share into the final solution: the DMSO fraction of the stock times the share of the final volume that stock takes up. Peptide suppliers set 1% (v/v) as the limit for DMSO in cell-based assays. JPT advises not exceeding it, and STEMCELL keeps the final concentration below it. At fixed target concentrations the total does not depend on the final volume, so making more solution does not dilute it. The planner warns you when cumulative DMSO is above 1%. DMSO stocks are thick (viscous) and dense, and a positive-displacement pipette transfers them more precisely (Rainin).
Keep copper complexes and free thiols apart until tested
Copper binds thiols (sulfur-containing groups on some amino acids) strongly and oxidizes them in vitro, that is, in a test tube (Eben and Imlay, 2023). A copper–tripeptide complex studied in the literature exchanges its copper quickly with other molecules of the same peptide and forms mixed complexes with glycine or histidine (Hureau et al., 2011). No study we found shows a copper–peptide complex oxidizing other peptides in a mixture. Even so, because copper complexes can hand copper over to competing ligands (other molecules that also bind it), do not combine one with free-thiol peptides or reducing agents without testing first. A strongly acidic stock can also compete for the copper: protons compete with the metal, and copper binding to these sequences sets in only around pH 3 to 4 (Gonzalez et al., 2018).
Oxidation-prone residues
JPT advises against DMSO for peptides containing methionine or cysteine, because it may oxidize the side chain, and suggests DMF (dimethylformamide) instead. Cys, Met and Trp are prone to oxidation in any case, a reaction with oxygen that alters the amino acid. It is faster during freeze–thaw cycles and at high pH (Hoofnagle et al., 2016), and Bachem notes that free cysteine thiols oxidize rapidly above pH 7. A free-thiol component can also exchange with another component’s disulfide, and the exchange speeds up as the pH rises (Nagy, 2013). As a precaution, the planner flags any Cys, Met or Trp component that comes from a DMSO stock.
Losses to the wall are unequal
When you dilute a mixture, its components are not lost to the container wall equally. In Thermo Scientific’s calibration mixture, three of the fifteen peptides gave a 20 to 60% lower signal, and a more variable one, when stored at the lower concentration. Keep every component of the most dilute solution above the range where such losses begin, and dilute from concentrated stocks close to the time the method runs.
These are rule-based checks that follow supplier guidance and methods papers. A mixture that raises none of them has not thereby been shown to be compatible.
How does the planner calculate a mixture?
Each component is its own small dilution into one shared final volume, and the diluent (the liquid you add) makes up whatever volume the stocks leave.
For each component, the transfer from its stock is v = c × V ÷ S, where c is the component’s target concentration in the final solution, V is the final volume and S is the stock concentration. The diluent is V minus the sum of the transfers. If you have met C1V1 = C2V2, this is the same rule, applied once per component.
The planner works in three modes:
- Stocks into an empty container. You give the final volume and, for each component, a stock concentration and a target. The planner returns every transfer and the diluent.
- Combining whole preparations. Each component is dissolved in full and everything is combined. Each final concentration is that component’s amount divided by the total volume, so the amounts you start with fix the ratio between components.
- Adding to an existing solution. The base solution already holds one component. The planner finds the final volume that brings it to its target, works out the transfer for each new component, and checks the total against the container.
A plan only works when the transfers fit inside the final volume: the sum of c ÷ S over all components must be 1 or less. Each term is the share of the final volume that one stock takes up. Every transfer scales with the final volume, but the shares do not, so changing the final volume never rescues a plan that does not fit.
When a plan does not fit, the planner does not simply say no. It names the components with the largest shares and offers two ways out: a more concentrated stock for one of them, up to the solubility ceiling you entered, or a lower target concentration. Leaving a component out of the mixture also reduces the sum.
The cumulative co-solvent follows the same rule. Each DMSO stock contributes its DMSO fraction times its share of the final volume, and at fixed targets that total is the same at any final volume.
The planner applies the concentration calculator’s checks to every transfer and to the total: a transfer below 2 µL is an error and one below 10 µL a caution, and a vial or tube filled above 90% of its capacity triggers a warning. These checks are there to catch trouble before you reach the bench.
- Transfer
- v = c × V ÷ S, for each component
- Diluent
- V − Σv
- Feasible when
- Σ(c ÷ S) ≤ 1, at any final volume
- Cumulative DMSO
- Σ(c ÷ S × the stock’s DMSO fraction), also independent of the final volume
Two worked plans: one that fits and one that does not
The same three-component arithmetic, twice: in the first plan the stocks are concentrated enough, and in the second they are not.
Plan 1 fits. Components A, B and C are each wanted at 100 µg/mL in 1.000 mL, from stocks at 5.000, 5.000 and 10.00 mg/mL. In µg/mL the stocks are 5000, 5000 and 10000. So A needs 100 × 1.000 ÷ 5000 = 0.02000 mL, B needs the same, and C needs 100 × 1.000 ÷ 10000 = 0.01000 mL. That is 20.0, 20.0 and 10.0 µL. The diluent is 1.000 mL less 50.0 µL, or 950.0 µL. The stocks take up 5% of the final volume, and no transfer is below the 10 µL caution threshold.
Plan 2 does not fit, and seeing why is useful. A, B and C are each wanted at 1.000 mg/mL in 1.000 mL, from stocks at 2.000, 2.000 and 5.000 mg/mL. The transfers would be 0.500, 0.500 and 0.200 mL, 1.200 mL in all, but the final volume is only 1.000 mL, which would leave −0.200 mL for the diluent. The shares are 0.5, 0.5 and 0.2, so Σ(c ÷ S) = 1.2, above 1. The planner names A and B, each half of the final volume, as the largest shares.
Making more solution does not help. At 2.000 mL, the same targets need 1.000, 1.000 and 0.400 mL of stock, 2.400 mL in all, so the plan is 20% over at any volume.
Changing the shares does help. With the A and B stocks at 4.000 mg/mL, if both peptides dissolve that far, the transfers become 0.250, 0.250 and 0.200 mL, with 0.300 mL of diluent. Lowering the A and B targets to 0.500 mg/mL gives the same volumes. Either change brings Σ(c ÷ S) to 0.7.
- Stock A20.0 µL
- Stock B20.0 µL
- Stock C10.0 µL
- Diluent950.0 µL; final solution 1.000 mL.
A pre-blended lyophilized mixture, or separate stocks?
Both give you a solution of several peptides. They differ in what you can choose, check and control.
One powder, one fixed ratio
A three- or four-component lyophilized (freeze-dried) mixture dissolves as one: every component goes into the same first solvent, which is a compromise when the components differ in charge. The ratio is fixed when the powder is made, and a single-component control needs each peptide from another source. Because one purity figure is not defined for a mixture, its certificate needs an identity finding and a peak-area reading for each component.
Each component dissolved on its own terms
Each peptide is dissolved in the first solvent its sequence suggests, checked for clarity on its own, and then combined at a ratio you choose and record. The same stocks supply the single-component controls. The trade-off is that every stock brings its solvent into the mixture, which is what the chemistry checks above are for.
Take the composition from the certificate
An informal peptide nickname can point to two different things: an acetylated fragment of about 889 g/mol without methionine, or a full 43-residue chain of 4963.51 g/mol with one methionine. They are different substances. The identity test on a certificate of analysis tells them apart, and the planner needs the molecular weight of the one actually present to report µM. The methionine also matters for the DMSO check, so it is worth getting right.
Common questions about multi-component peptide solutions.
How do I prepare an equal-mass or an equimolar peptide pool?
Start by choosing the basis. An equal-mass pool has the same µg/mL of every peptide, so the molar amounts differ with molecular weight. An equimolar pool has the same µM of every peptide, and needs each component’s molecular weight and net amount. Commercial pools are specified per peptide, for example 25 µg or 15 nmol of each. Enter each component’s stock concentration and target in the planner; if you enter molecular weights, it reports µM beside mg/mL. For cell-based assays, keep the cumulative DMSO below 1% (v/v): a 1:100 dilution of a stock in 16% DMSO carries 0.16%.
How do I prepare a mixed calibration standard from stocks at different concentrations?
Treat each component as its own dilution into one shared final volume: the transfer is c × V ÷ S for each, and the diluent makes up the rest. For example, with stocks at 5.000, 5.000 and 0.500 mg/mL, and 100.0 µg/mL of each in a 10 mL volumetric flask, the transfers are 0.200, 0.200 and 2.000 mL, and the diluent, 7.600 mL, brings the flask to the mark. Use a fresh, pre-rinsed tip for each transfer, dispense into the diluent, and prepare the standard from concentrated stocks close to the time the method runs.
Why does a larger final volume not fix a plan that does not fit?
Because every transfer scales with the final volume. Each stock takes up the fraction c ÷ S of the final volume whatever that volume is, so fractions that add up to more than 1 at 1 mL still add up to more than 1 at 10 mL. Only a more concentrated stock (within its solubility), a lower target or fewer components changes the sum. The cumulative DMSO percentage behaves the same way.
Do peptides in one solution interact with each other?
They can, and for the peptides most often combined we found no published study of which ones do. That is a good reason for care. A free-thiol peptide can exchange with another component’s disulfide, faster as the pH rises, and free cysteine oxidizes rapidly above pH 7. A copper complex can hand its copper to competing ligands, and copper oxidizes free thiols in vitro, so do not combine a copper complex with free-thiol peptides or reducing agents without testing. Opposite charges matter mainly through pH: when isoelectric points are far apart, the shared pH can sit near one component’s solubility minimum. Verify the prepared mixture by HPLC or MS.
Can two separately prepared stock solutions be combined?
As arithmetic, yes. Combining stock solutions into a fixed final volume dilutes each component by the ratio of its own volume to the total. Two 1.000 mL solutions at 2.000 mg/mL, combined with no further diluent, give 1.000 mg/mL of each component in 2.000 mL. The chemistry is the open question. Check the solvents and pH of both stocks against the planner’s flags first, and expect the less stable component to set how long the mixture can be kept.
How long can a mixed peptide solution be kept?
There is no general answer, and we would rather say so than guess. Peptide solutions keep less well than powder, especially when a sequence contains Asn, Gln, Cys, Met or Trp, and in a mixture the least stable component sets the limit. We found no published stability data for the combinations most often sold pre-blended. Keep each component as its own concentrated stock, prepare the mixture close to the time the method runs, and verify it by HPLC or MS if it must be stored.