Evaluation of peptide concentration on stability of signal over time. The peak areas (normalized to time 0) from 2 representative peptides were plotted vs autosampler storage time (h) to show that storing the peptides at higher concentration can minimize the loss of peptide signals, presumably attributed to adsorption of the peptides to vials. For details, see online Supplemental Materials and Methods, which accompanies the online version of this article at  

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... or dimethyl- formamide can be used to efficiently solubilize peptides with a high percentage of hydrophobic residues ( Ͼ 50% Ala, Val, Leu, Ile, Met, Phe, Trp, Pro) and Ͻ 25% charged residues. Before reconstituting peptides, lyophilized powder should be brought to room temperature in a desiccator to avoid water absorption in the unused peptide, thus minimizing variations in concentration of lyophilized aliquots. If reconstituting a peptide for the first time, and whenever possible, a small amount of the peptide should be reconstituted before committing the entire lot by weighing out a small aliquot. As discussed above, the pH is an important parameter for peptide solubilization. Ini- tial reconstitution is best performed in water by adjusting the pH based on the primary amino acid sequence, with a small amount of organic solvent added to aid solubilization. Buffers such as PBS should not be used for reconstitution, because salts hinder solubility. If salt solutions are desired for the final formulation, they are best added once the peptides are fully solubilized. Peptides should initially be reconstituted at a concentration that is higher than the desired final working concentration (typically 10 –1000 times more concentrated; see Table 7 for specific recommendations). Solutions of completely solubilized peptides are completely clear and are devoid of any flecks or cloudiness. Solubili- zation can be confirmed by light scattering analysis or by comparing absorbance in a series of dilutions with and without centrifugation to pellet undissolved material. A general recommended starting point for a reconstitution solution is 5% acetonitrile with 0.1%–1% formic acid. The inclusion of organic solvent and acid in the reconstitution solution not only aids solubility, but also serves to retard microbial growth (biologically active buffers should contain 0.1% sodium azide to prevent microbial growth). If this reconstitution solution is not successful in completely solubilizing the peptide, the amount of organic solvent can be increased or the organic solvent can be altered (e.g., methanol instead of acetonitrile). If increasing organic solvent is not effective in solubilizing the peptide, the pH can be adjusted by addition of acid ( Յ 1% formic acid or trifluoroacetic acid) or by use of 1% ammonium bi- carbonate, 1% N , N -diisopropylethylamine, or ammonium hydroxide. Another option is to redry the peptide and redissolve it in DMSO. Variable recovery because of nonspecific adsorption is 1 of the major consequences of improper handling of peptides and can lead to imprecision and bias (i.e., loss of peptide to surfaces or contamination/carryover). The extent of nonspecific peptide adsorption to the walls of peptide storage vessels, pipette tips, autosampler vials, and HPLC components varies on the basis of the primary sequence, the materials used, and the concentration of the peptide solution. Complete characterization of peptide stability includes the evaluation of losses due to adsorption in all steps of the analytical method. This can be accomplished by several experimental designs, including measuring peptide amounts in serial dilutions by UV absorbance (e.g., to evaluate potential loss in tubes and/or pipette tips) or repeated injections by LC-MS (e.g. to evaluate potential loss or carryover in vials and the HPLC system). The use of carrier or chaperone molecules can minimize adsorption effects for particularly difficult peptides (56 ) ; however, choice of a suitable carrier is highly dependent on the peptide sequence, the analytical method, and the desired matrix for analysis. Thus, there is currently no consensus related to the best carrier molecules or the optimum concentration for use with peptide internal standard and calibrators. When evaluat- ing carrier molecules, caution should be taken to choose components that do not interfere with detection of the target peptide or excessively contribute to sample complexity or instrument contamination. The relative loss of peptides by nonspecific adsorption in low-concentration solutions is greater than in more concentrated solutions because of the limited bind- ing capacity of the wetted solid surface area (57 ) . To demonstrate the loss of peptides in solution and the effect of storage concentration, 2 peptide mixtures (200 and 1000 fmol/ ␮ L) were prepared in nondeactivated glass vials and analyzed by injecting 1 ␮ L of each sample each hour for 15 h. Of the 50 peptide targets in each mixture, 48 and 50 peptides were detected in the 200- and 1000- fmol/ ␮ L samples, respectively. Nine and 0 peptides, respectively, showed noticeable signal decay over time under the 2 conditions. This effect can be seen by plotting total peak areas of 2 representative peptide sequences, YLGYLEQLLR [Sequence Specific Retention Calculator (SSRC) relative hydrophobicity 41.55] and IYEGSI- LEVDCDILIPAASEK (SSRC relative hydrophobicity 43.98), both of which are quite hydrophobic (Figure 3). In contrast to the 200-fmol/ ␮ L sample, all peptides in the 1000-fmol/ ␮ L mixture showed constant signals over the time period analyzed, consistent with improved stability and reduced adsorption at higher concentration. Nonspecific adsorption contributes to carryover, which increases variability and bias due to residual signal in sample runs (58 ) . Carryover in sample preparation can originate from reusing pipette tips to transfer peptide solutions between vials or in dispensing aliquots. Carryover in sample preparation or analysis can negatively affect results through ion suppression of low-abundance peptides (when coelution occurs with high-abundance carryover from the previous run or sample) or by producing a false positive in sample analysis by the detection of contaminating analyte peptide. One can determine the extent of nonspecific adsorption by transferring a solution of the analyte sequentially from 1 vial to another and analyzing a small aliquot after each transfer step to assess for losses (59 ) . Despite the diverse physicochemical properties of peptides, various strategies can be generically applied to reduce adsorption and cross-contamination phe- nomena (56, 60 ) leading to carryover. When preparing dilution series, one should never reuse pipette tips, to avoid cross-contamination. Pipette tips should be pre- rinsed several times with the peptide solution before aspirating the final volume. To minimize nonspecific adsorption to the walls of storage vessels, standards of peptides should be added directly to the diluent fluid instead of the sides of the tubes or vials. Finally, peptide adsorption also contributes to carryover in chromatographic systems through incomplete removal of analyte from the analytical system from the previous injection (e.g., insufficient wash of the injection valve or syringe of the autosampler). Chromatographic carryover can be evaluated by injecting a blank sample after a sample or calibrator. Complete system wash runs (e.g., rinsing all HPLC components, including autosampler, delay volumes, and columns) can be used to reduce or eliminate carryover with a series of different elution buffers and solvents. It should be noted that some peptides, especially those containing hydrophobic residues, can be retained on HPLC columns despite the use of high concentrations of organic solvents when washing. Most HPLC column manufacturers have published methods for cleaning the HPLC flow-path and columns. Different types of vials can introduce significant variability in LC-MS analyses (61 ) . The interaction of peptides with various surfaces is greatly influenced by the specific side chains of the amino acids of the peptide. Glass and polypropylene are the materials most commonly used to manufacture vials, inserts, and plates. Although a single type of vial might not be optimal in terms of minimizing the nonspecific interaction of all the peptides in an analytical mixture, basic amino acids can form electrostatic interactions with the residual silanol groups on glass vials, and nonpolar amino acids can interact with the hydrophobic surface of polypropylene vials (62 ) . To minimize these adverse interactions, several manufacturers of chromatography consumables offer silanized glass vials in which the silanol groups have been chemically inactivated. Similarly, polypropylene vials with modified plastic surfaces are commercially available. To demonstrate the variability that can arise from various container materials, we investigated the signal from repeated injections of a digested protein sample stored in 3 types of autosampler sample vials: nondeactivated glass, deactivated glass, and polypropylene vials. Peptide stability was tested by performing 15 repeated LC-MS/MS analyses of the 50-fmol/ ␮ L sample each hour for 15 h. We manually assessed the signal intensities of the replicate runs for each peptide to determine the amount of signal enhancement or decay. The results are summarized in Figure 4. Peptides were categorized as stable, slow decay, or fast decay, with cutoffs of Ͻ 5%, 5%–50%, or Ͼ 50% peptide loss on the basis of signal intensity over the 15 h. We found that all 3 vial types enabled the recovery of 43 peptides, which accounted for 86% of the monitored peptides. Twenty-nine of the detected peptides were very stable across all analyses for all vials. In this study, the polypropylene vial outperformed the 2 glass vials, as only 1 “unstable” peptide with signif- icantly lower recovery was detected, whereas 13 and 14 unstable peptides were detected in nondeactivated and deactivated glass vials, respectively. To demonstrate the effects of freeze–thaw on peptide stability, we compared the signal intensity observed when injecting a peptide mixture stored at 4 °C, a sample undergoing a single freeze–thaw, and a sample undergoing multiple (n ϭ 10) freeze–thaw cycles. Twelve 1-pmol/ ␮ L sample aliquots prepared in solution (3% acetonitrile, 0.1% formic acid, ...