Dual-Purpose Polymer Labels for Fluorescent and Mass Cytometric

Apr 10, 2013 - ... available by participants in Crossref's Cited-by Linking service. ... in the quantitative analysis of biomolecules with exogenous t...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/Biomac

Dual-Purpose Polymer Labels for Fluorescent and Mass Cytometric Affinity Bioassays Daniel Majonis,†,‡ Olga Ornatsky,‡ Dirk Weinrich,† and Mitchell A. Winnik*,† †

Department of Chemistry, University of Toronto, 80 St. George Street, Toronto, Ontario, Canada, M5S 3H6 DVS Sciences, 70 Esna Park Drive, Unit 12, Markham, Ontario, Canada, L3R 6E7

‡

S Supporting Information *

ABSTRACT: We describe the synthesis and characterization of a family of poly(N-alkylacrylamide) polymers carrying 2−6 fluorescent dye molecules, ∼70 pendant DTPA (diethylenetriaminepentaacetic acid) groups, and an orthogonal maleimide end-group for covalent attachment to an antibody (Ab). These dual-purpose labels were designed for use in multiplexed immunoassays based on both mass cytometry and fluorescent flow cytometry. A challenge in the polymer synthesis was finding conditions for attaching a sufficient number of dye molecules to each polymer chain. Although attachment of a terminal maleimide to the polymers was not as efficient as anticipated, the end-functional polymers were still effective in labeling Abs. Secondary goat antimouse IgG was labeled with the four dual-label polymers as well as a control polymer, and while the resultant antibody-polymer conjugates showed positive performance in mass cytometric and fluorescent assays, some trials showed problems such as low signal and nonspecific adsorption. Four primary antibody conjugates were prepared and used to stain cells in 4-plex assays. The results of both primary assays are bittersweet in that the CD3-FITC and CD45-DyLight 649 conjugates performed well, while the CD13-DyLight 405 and the CD38-DyLight 549 conjugates did not.

■

INTRODUCTION Mass cytometry is a new technique designed for highly multiplexed cell-by-cell immunoassays.1 In this technique, various antibodies are labeled with multiple copies (>100) of a different metal isotope. After treating a cell suspension with a cocktail of these antibodies, the cells are injected individually but stochastically into the plasma torch of an inductively coupled plasma mass spectrometer (ICP-MS) equipped with time-of-flight detection. In the torch, the cells are vaporized, atomized, and ionized to generate metal ions whose abundance is measured quantitatively.2 The basic principle of cell staining with a cocktail of labeled antibodies followed by cell-by-cell analysis is similar to that of flow cytometry. Flow cytometry is an established technology and benefits from a wide range of commercially available reagents and simpler instrumentation. Fluorescence detection, however, has a number of limitations. Fluorescent dyes have relatively broad emission spectra, which limits the number of dyes that can be detected in a single experiment. On a routine basis, 4-plex assays are common, but extending the technique to 15+ dyes remains a serious challenge. The dynamic range is only 2 to 3 orders of magnitude, compensation between different fluorophores is required, and cell autofluorescence can be problematic. The features of mass cytometry that make it particularly attractive for biomarker detection is that multiplexing capabilities are limited only by the availability of metal isotopes with masses in the range of 100 to 200 amu. Currently 31 metal isotopes, primarily lanthanide (Ln) isotopes, are commercially © XXXX American Chemical Society

available. This high multiplexity is particularly useful when a researcher’s cell analysis requires many surface or intracellular cell markers be monitored at once. For example, Bendall and co-workers monitored a total of 49 markers by staining cells with one of two sets of labeled antibodies; the first set monitored 13 core cell-surface markers with an additional 18 surface markers, and the second set monitored the same 13 core markers as well as 18 intracellular epitopes.3 In addition, the technique of mass cytometry has a large dynamic range, and background signal is normally negligible. Furthermore, it does not require compensation between signals, and mass spectrometry detection is quantitative.3 One of the shortcomings of mass cytometry is that the cells detected and analyzed are vaporized and cannot be sorted and collected, as in fluorescence activated cell sorting (FACS). Mass cytometry is a new technique while FACS is more established; thus it is important to carry out comparative studies using both techniques for biomarker detection. While some experiments of this type have been reported,3 we are interested in developing dye-labeled metal chelating polymers for attachment to antibodies that will serve as dual-purpose staining reagents. Antibodies labeled with these polymers can be employed for cell staining and analyzed by both techniques. Received: February 1, 2013 Revised: March 21, 2013

A

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

nide signal to the cells. However, signal was not as high as that obtained with the X8 conjugate, and nonspecific adsorption to the EL4 cells was observed for two of the fluorescent conjugates. In another experiment, GAM was conjugated to the FITC-labeled polymer and was used in a sandwich assay to stain Ramos cells that were previously stained with primary CD45 antibody. Analysis by mass cytometry as well as by FACS separately demonstrated that the cells were stained with both lanthanide and fluorescent markers. Next, four primary labeled antibodies were created with CD3, CD13, CD38, and CD45 and the four different fluorescent polymers and subsequently used to stain KG1a and Jurkat cells. Independent analysis by mass cytometry and FACS demonstrated that the FITC and DyLight 649 antibody tags performed well, but the other two tags did not. Modification of the polymer with the other two dyes followed by attachment to the antibodies appears to interfere with the cell-staining capabilities of the antibodies. Further experiments are required to determine why the DyLight 405 and 549 antibody tags failed to perform satisfactorily in the bioassays.

Dual-purpose tags of this sort will enable a number of direct applications. First, FACS and mass cytometry assays performed independently in parallel will help to further validate the technique of mass cytometry. Second, dual-labeled tags will enable a researcher to perform a rapid assay with FACS, followed by a higher multiplexity mass cytometric assay. Third, the dual tags will allow one to sort samples by FACS4,5 using dye markers prior to further detailed multiparametric analysis by mass cytometry. This approach is useful in situations where one is interested in a small cell subset present in a heterogeneous population of cells. Appropriate steps for achieving this goal would include staining cells with 1−36,7 dual-purpose antibody tags that recognize biomarkers characteristic of the cell subset of interest, sorting the cells by FACS, staining the isolated cells with additional, but nonfluorescent, metal-labeled antibody tags, and finally analyzing the cells by mass cytometry. One approach to create a dual-purpose antibody tag is to functionalize an antibody separately with both commercially available reactive fluorescent dyes and metal-chelating moeties.8,9 However, as a general rule, it is best to limit the degree of antibody modification; if an antibody is overmodified, it will lose specificity for its target.10 The most straightforward approach is to create an all-in-one probe that contains both fluorescent dye molecules and metal-chelating groups.11 In our previous publications describing metal-chelating polymers for mass cytometry, the goal was to create polymers with a narrow length distribution and with a pendant ligand such as DTPA on every repeat unit of the polymer chain, and a maleimide at the chain end for antibody attachment.2,12 In this work, our aim is to retain those features while also including 1− 5 pendant fluorescent dye molecules per chain. One challenge here in the synthetic design was to add the fluorescent dye(s) in an efficient manner and at a late stage of the synthesis. This is necessitated by the high price of modern, reactive fluorescent dyes. It also significantly simplifies the synthesis because one precursor polymer can be used to create polymers with different dyes. Another challenge is to maintain the same high DTPA functionality obtained with the earlier nonfluorescent polymers. In the work described here, we begin our synthesis with a poly(N-alkylacrylamide) identical to that described previously, a polymer that has a pendant primary amino group on each of the approximately 80 repeat units.2 Our strategy was to add short, protected PEG amino spacers to about 12% of the units, add DTPA groups to the remaining 88%, then deprotect and functionalize the PEG amino spacers with amine-reactive fluorescent dyes. We chose four dyes: FITC as a relatively inexpensive reagent to develop the chemistry, and DyLight 405, 549, and 649 to include additional colors in the fluorescence assay. The product polymers have 2−6 dye molecules per chain and nearly the same lanthanide binding capacity as the nonfluorescent polymer. The polymers were functionalized at one chain end with a maleimide linker. Then the polymers were loaded with stable lanthanide isotopes, and used to create dualpurpose antibody tags. The first labeled antibodies to be created were conjugates of all four fluorescent polymers, as well as of a commercially available control X8 polymer, with goat antimouse IgG (GAM). These GAM conjugates were used in a sandwich assay to stain KG1a and EL4 cells that previously were stained with primary CD34 antibody. Analysis by mass cytometry showed that the fluorescent polymer−antibody conjugates did impart lantha-

■

EXPERIMENTAL SECTION

Polymer Synthesis. Details of polymer synthesis are presented in the Supporting Information. Instrumentation and Characterization. 1H NMR (400 or 500 MHz) spectra were recorded on a Varian Hg 400, Varian 400, or Varian Unity 500 spectrometer with a 45° pulse width and at a temperature of 25 °C. All samples were dissolved in D2O, with chemical shifts referenced to the HDO peak at 4.77 ppm.13 Acquisition parameters included 512−768 transients and a delay time of 10 s. The nominal molecular weights and polydispersities of all anionic, water-soluble samples were measured with a Viscotek size exclusion chromatograph (SEC) equipped with a Viscotek VE3210 UV/vis detector, VE3580 refractive index detector, and Viscotek ViscoGEL G4000PWXL and G2500PWXL columns (kept at 30 °C). The flow rate was maintained at 1.0 mL/min using a Viscotek VE1122 Solvent Delivery System and VE7510 SEC Degasser. An eluent of 0.2 M KNO3, 200 ppm NaN3, and 25 mM pH 8.5 phosphate buffer was used. The system was calibrated with poly(methacrylic acid) standards. Samples were dissolved in eluent prior to injection. UV/vis spectra of the dye-labeled polymers were collected on a Perkin-Elmer Lambda 35 UV/vis spectrometer. Polymer samples (ca. 0.2 mg) were accurately weighed on a Mettler Toledo MX5 microbalance, transferred to 20 mL scintillation vials with Teflon tape-wrapped threads, then dissolved in a weighed amount of phosphate buffer (200 mM, pH 8.00). The polymers were assumed to carry 2.9 H2O and 2.2 Na+ per DTPA unit.12 This assumption and DPn values obtained from the 1H NMR analysis were combined with the results of the UV/vis measurement to calculate the number of dye molecules per chain. Excitation and emission fluorescence spectra of the Yb-loaded maleimide polymers were collected on a Jobin Yvon Horiba FL3−22 Fluorolog. Solutions were prepared in phosphate buffer (200 mM, pH 8.00) at a concentration where the primary peak of the fluorescent dye had an absorbance of 0.10. The metal content of a given sample was determined by traditional inductively coupled plasma-mass spectrometry (ICP-MS). An aliquot of the solution prepared for UV/vis spectroscopy was diluted by a factor of 4 to 10 with 2% HCl, after which a 5 μL aliquot of that solution was diluted to 5000 μL with 2% HCl. Diluted samples were analyzed on a Perkin-Elmer SCIEX Elan 9000 ICP-MS equipped with an autosampler. The ICP-MS signal was converted to ppb through the concurrent analysis of separate 1 ppb metal standards. Separately, a theoretical metal concentration that would be obtained on analysis of a fully metal-loaded polymer was calculated. This theoretical concentration was calculated from the polymer mass concentration and by B

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

assuming that each DTPA unit carried one lanthanide, one Na+, and 2.9 H2O.12 Finally, the number of metal atoms per chain was calculated by dividing the experimentally determined metal concentration by the theoretical metal concentration. Biological Experiments. Materials. Goat antimouse (GAM) was obtained from Pierce Biotechnology, CD3, CD13, CD34, and CD38 antibodies were obtained from Biolegend, and CD45 antibody was obtained from eBioscience. Ramos, KG1a, and Jurkat cells were obtained from ATCC (American Type Culture Collection, Manassas, VA). Antibody Labeling with Metal-Chelating Polymers. Metal-labeled antibodies were prepared as follows. In advance, 0.2 mg of maleimideterminated polymer was loaded with lanthanide ions, dried on an Eppendorf Vacufuge Plus, vacuum-packed, and stored in a −30 °C freezer, as described in the Supporting Information. The day of the antibody labeling, an antibody at 1 mg/mL in 150 mM sodium phosphate buffer, pH 7.2, and in the absence of bovine serum albumin (BSA) or gelatin was subjected to mild reduction by TCEP (tris(2carboxyethyl)phosphine) to convert the disulfides in the Fc fragment to thiols. The reduction and subsequent antibody-polymer conjugation steps were performed in 0.5 mL, 50K MWCO centrifugal devices (Millipore Amicon Ultra). The maleimide group of the purified, lanthanide-loaded metal-chelating polymer was bound to the thiol groups of the partially reduced antibody in Tris-buffered saline (TBS, 25 mM Tris, 150 mM NaCl, 2 mM KCl, pH 7.4). The metal-tagged antibody was washed several times in EDTA-free TBS and stored at +4 °C. When loaded with lanthanide ions, these polymers are resistant to leaching, and we have never observed any exchange of lanthanide ions between differently tagged antibodies when deployed in a multiple antibody staining cocktail.14 Metal Atoms Per Antibody. The number of metal atoms carried by each antibody was determined by a combination of UV/vis spectroscopy and ICP-MS analysis.2 The metal-labeled antibody was resuspended in TBS. Protein concentration was measured using a Nanodrop ND-1000 UV/vis spectrometer (Thermo Fisher Scientific, U.S.A.). An aliquot of labeled antibody was diluted 1:100000 in 2% HCl, and 0.1 mL was analyzed by ICP-MS using a Perkin-Elmer SCIEX Elan 9000 ICP-MS equipped with an autosampler. The ICPMS signal was converted to ppb through the concurrent analysis of separate 1 ppb metal standards. This data allowed the determination of the number of lanthanide atoms per mL. Combined with the Nanodrop data, which estimated the number of antibody molecules per mL, the mean number of lanthanide atoms per antibody molecule could be calculated. Fluorescent and Mass Cytometric Assays. All antibody staining steps utilized 50 μL of staining antibody solution per cell pellet of 1 × 106 cells. To test the four polymers in the first proof of concept assay, four different polymer-GAM conjugates were prepared using P(12% PEGAmino)(88%DTPA)( 169 Tm)(DyL405)-maleimide, P(12% PEGAmino)(88%DTPA)( 1 5 2 Sm)(FITC)-maleimide, P(12% PEGAmino)(88%DTPA)(165Ho)(DyL549)-maleimide, and P(12% PEGAmino)(88%DTPA)(159Tb)(DyL649)-maleimide. As a positive control, a polymer-GAM conjugate was also prepared with an aliquot of commercially available (DVS Sciences) X8 metal-chelating polymer, loaded with 159Tb. KG1a and EL4 cells were separately stained with primary CD34 followed by one of the secondary GAM-tags. Washed cells were fixed in 3.7% formaldehyde and counterstained with an Ir intercalator15 for nucleated cell identification. To test the FITC-labeled polymer in the second and third proof of concept assays, a polymer-antibody conjugate was prepared with P(12%PEGAmino)(88%DTPA)(172Yb)(FITC)-Maleimide and GAM. Ramos cells were stained with primary CD45 followed by the secondary GAM-tag. As above, washed cells were fixed in 3.7% formaldehyde and counterstained with an Ir intercalator prior to analysis. To test all four polymers in the mass cytometry and FACS tetraplex assays, polymer-antibody conjugates were prepared with the four dyelabeled polymers and CD3, CD13, CD38, and CD45, as listed in Table 1. KG1a and Jurkat cells were stained with six titration solutions. Each

Table 1. Dye Functionalization, Dye Characteristics, and Remaining Lanthanum for P(12%PEGAmino)(88% DTPA)(DYE)-Disulfide Polymer Samples, and Maleimide Functionality for (12%PEGAmino)(88%DTPA)(DYE)Maleimide Polymer Samples

sample FITC DyLight 405 DyLight 549 DyLight 649

dye equivalents used per chaina

dye per chainb

ε (M−1 cm−1) per dye moleculec

remaining La3+ per chaind

maleimide groups per chaine

30 8.5

3.4 6.2

88000 30000

∼0 0.3

0.21

6.9

3.9

150000

0.4

0.46

6.7

2.6

250000

0.2

0.40

a

Equivalents of reactive dye used in each dye labeling reaction. Resultant number of dye molecules per chain. The maximum possible number is 9. cExtinction coefficient values are taken from the literature for FITC16 and from manufacturer specifications for the DyLight dyes.22 dRemaining average number of lanthanum ions retained per chain, as determined by solution ICP-MS. eThis value could not be determined by 1H NMR for FITC. b

staining solution contained a different concentration of all four antibody tags, as shown in Table S4 in the Supporting Information. As above, washed cells were fixed in 3.7% formaldehyde and counterstained with an Ir intercalator prior to analysis. One batch of labeled cells were analyzed by mass cytometry,1 using a CyTOF instrument from DVS Sciences, Markham, Ontario. Mass cytometry is a real-time analytical technique whereby cells or particles are individually introduced into an inductively coupled plasma, and each resultant ion-cloud is analyzed by time-of-flight mass spectrometry. Dual-counting, the combination of digital counting and analog modes of ion detection, allows a much wider range of ion signal (simultaneous detection of very small and very large signals). Another batch of labeled cells were also analyzed by FACS, on a BD Biosciences LSRII instrument. DyL405, FITC, DyL549, and DyL649 fluorescent signals were monitored with the laser and filter setups of the instrument intended for Pacific Blue (excitation 405 nm, emission peak max 440 nm with bandwidth at half-maximum range of 40 nm “440/40 nm”), FITC (excitation 488 nm, emission 530/30 nm), phycoerythrin (excitation 532 nm, emission 575/26 nm), and Alexa fluor 647 (excitation 633 nm, emission 660/20 nm), respectively. Unstained Ramos or KG1a cells were run as a negative control to determine background fluorescence signal. Mass cytometry and FACS data were collected in FCS 3.0 format and processed by FlowJo (Tree Star Inc., Ashland, OR) software.

■

RESULTS AND DISCUSSION Polymer Synthesis. We began the synthesis with a DPn = 79 amino polymer-disulfide (1). As described previously,2 the synthesis of this polymer began with the reversible addition− fragmentation chain transfer (RAFT) synthesis of poly(tertbutyl acrylate). Through a number of postpolymerization modifications, we obtained an acrylamide polymer with an average of 79 primary amines and an end-functional thiol group, protected as a polymeric disulfide. We have previously shown that2,12 the pendant primary amines of this polymer can be quantitatively functionalized with DTPA. Then the disulfide was reduced to a pair of thiol groups, which, in turn, were reacted with a maleimide. Here we describe the selective introduction of two different pendant groups to the polyacrylamide: DTPA groups to bind lanthanide ions, and a much smaller fraction of PEG spacer chains to which fluorescent dyes can be attached. These C

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

Scheme 1. Synthesis of Fluorescent Metal-Chelating Polymersa

a

HEPES = 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid buffer, DMF = dimethylformamide, EDC = 3-(ethyliminomethyleneamino)N,N-dimethyl-propan-1-amine, NHS = N-hydroxysuccinimide, DTPA = diethylenetriaminepentaacetic acid, DMTMM = 4-(4,6-dimethoxy-1,3,5triazin-2-yl)-4-methylmorpholinium chloride, TFA = trifluoroacetic acid, PB = phosphate buffer, DMSO = dimethyl sulfoxide.

serves two functions. The first is to separate the dye from the polymer backbone. The second is to add a small number of protected amino groups before the reaction in which DTPA groups are added. This strategy allowed us to react dyes with these amino groups in the penultimate synthetic step, minimizing the number of reactions to which the dye is exposed. Reaction conditions were optimized with FITC as the amine-reactive fluorescent dye. The first step in the synthesis was the EDC/NHS-mediated coupling of N-Boc-N′-succinyl-4,7,10-trioxa-1,13-tridecanediamine to polymer 1. The mean degree of modification was 9 units per 79 unit polymer chain, as determined from the 1H and gCOSY (gradient correlation spectroscopy) NMR spectra presented in Figures S1 and S2 of the Supporting Information, respectively. We assume that the resultant polymer is a random copolymer. Next, the remaining primary amino groups were functionalized with DTPA in the presence of 4-(4,6-dimethoxy1,3,5-triazin-2-yl)-4-methylmorpholinium chloride (DMTMM) as the coupling agent, as we have reported previously.2 Following this step, the polymer was treated with 50% TFA in water for 2 h at room temperature to remove the BOC protecting group from the terminal amines of the short PEG linker. Deprotection was confirmed by the complete disappearance of the BOC tert-butyl peak in the 1H NMR spectrum of the product (Figure S3). In Scheme 1, we refer to the disulfide polymer partially functionalized with the BOC-protected amine as 2, and the specific polymer used for subsequent transformations as P(12% PEGAminoBoc)-disulfide, where the 12% refers to the fraction of pendant groups carrying the PEG spacer, as determined by 1 H NMR. Similarly, we refer to the copolymer containing both

pendant group transformations are depicted in Scheme 1. In the first step, a fraction of the amino groups of 1 were reacted with N-Boc-N′-succinyl-4,7,10-trioxa-1,13-tridecanediamine. The extent of conversion could be controlled by variation of the concentration of reactants and monitored by 1H NMR. In the second step, the remaining amino groups were reacted with a large excess of DTPA to form a pendant DTPAmonomamide. After deprotection of the BOC-amine at the end of the PEG spacers, an amine-reactive dye was attached. There are some subtle features of this reaction that will be discussed in more detail below. For example, the reaction with the dye was inefficient unless the DTPA groups were first saturated with La3+ ions, which were removed by ion exchange after the dye coupling reaction. In the final step of the reaction, the disulfide bond was reduced, followed by reaction of a large excess of the bismaleimide shown in the scheme. In the following paragraphs, we will examine each of these reactions in the context of our experimental design. The PEG spacer was chosen to separate the fluorescent dye from the polymer backbone to minimize the polyelectrolyte effect of the polymer on the degree of ionization of the dyes. Fluorescein, for example, has pKa values of 2.2, 4.4, and 6.3, and exhibits efficient fluorescence only when fully depronated.16 These pKa values are raised by an anionic microenvironment, resulting in poor fluorescence at neutral pH.17 The lanthanideloaded DTPA pendant groups will be anionic at neutral pH;18 thus our concern was that this anionic microenvironment may have a similar effect on fluorescein. This problem can be avoided through the use of fluorescein derivatives with electron-withdrawing groups,19 but FITC is attractive because of its availability and reasonable cost. The short PEG spacer D

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

almost all of the lanthanum from the polymers, as monitored by ICP-MS analysis. Aqueous SEC analysis of the product showed the previous additional higher molecular weight shoulder had disappeared, and the main polymer peak shifted almost completely back to the original molecular weight of the P(12%PEGAmino)(88%DTPA)-disulfide (Figure S4). These optimized reaction conditions were then applied to three NHS-ester DyLight dyes: 405, 549, and 649. Lower equivalents of reactive dye were used, but similar levels of functionalization were obtained. Table 1 collects data on dye functionalization, dye characteristics, and remaining lanthanum ions per chain. The normalized absorption spectra of the P(12%PEGAmino)(88%DTPA)(DYE)-disulfide polymers are presented in Figure S5. Attachment of the Bismaleimide Linker. The final steps in the polymer synthesis involved reduction of the disulfide bond with DTT, followed by the addition of a bismaleimide linker molecule. As in our previous reports,2,12 the outcome of this reaction was measured by 1H NMR by comparing the 5H signal of the phenyl end-group to the 2H signal of the maleimide group. Partial 1H NMR (D2O) spectra of all four polymer samples are presented in Figures S6−S9. We could not determine the maleimide yield of the FITC-labeled polymer due to overlapping peaks. According to the 1H NMR analyses, the DyLight 405, 549, and 649 polymer samples had 0.21, 0.46, and 0.40 maleimide groups, respectively, per chain. This is a lower level of maleimide incorporation than we reported previously2,12 for analogous polymers based on the polyacrylamide backbone, protected as a dimeric disulfide during pendant group transformations. In 2010,2 we reported 0.8 to 0.9 maleimides per polymer for a polymer of similar chain length with a DTPA on every repeat unit. Reinvestigation of this polymer (see Supporting Information for more details) led to a somewhat smaller value, 0.68 maleimides per polymer. The lower value appears to be a consequence of a more stringent purification protocol to remove bismaleimide noncovalently associated with the polymer. In 2011,12 we showed that a series of polymers of identical chain length (DPn ≈ 80), each with a different polyaminocarboxylate chelator (ethylenediaminetetraacetic acid, EDTA; DTPA, triethylenetetraaminehexaacetic acid, TTHA, and 1,4,7,10-tetraazacyclododecane-,4,7,10-tetraacetic acid, DOTA), all yielded quantitative end-labeling with N-(5-fluoresceinyl)maleimide, following reduction of the disulfide with DTT. These results show that the reduction step with DTT is not problematic. It should not be surprising that N-(5-fluoresceinyl)maleimide gives a higher yield of adduct with the end-group thiol than the bismaleimide. Friedman and Wall23 showed that rate constants for the formation of Michael adducts increase with the electronwithdrawing and resonance stabilizing nature of the substituents conjugated to the double bond of the Michael acceptor. Subsequent publications24,25 showed that the rate constant for the reaction of glutathione with N-(5fluoresceinyl)maleimide at pH 7.4 is about 7 times larger than the (extrapolated) rate constant for the reaction of glutathione with N-ethylmaleimide at pH 7.3. This difference in reactivity provides a reasonable rationale for the difference in results between 2,2′-(ethylenedioxy)bis(ethylmaleimide) and N-(5-fluoresceinyl)maleimide. Interestingly, the bismaleimide reaction of the EDTA polymer,12 utilizing the updated wash procedure, yielded 0.90 maleimide groups per chain. Thus, the success of a maleimide reaction depends both on the pendant

PEG and DTPA groups as 3, and the specific example examined in this study, after saturation with LaCl3, as P(12% PEGAminoBoc)(88%DTPA)(La)-disulfide. Polymer 4 refers to all of the dye-labeled polymers with a terminal maleimide. Optimizing the Attachment of Fluorescent Dyes. Initial attempts to functionalize P(12%PEGAmino)(88%DTPA)Disulfide by direct reaction with FITC met with limited success. These experiments were performed in phosphate buffer (200 mM, pH 7.50 or 8.50) with DMSO (33% v/v) and 24 equiv of FITC per chain. Analysis of the absorption spectrum of the product suggested that the dye was attached to only 2−3% of the PEG-spacer amino groups (only 0.2−0.3 FITC per chain). This level of dye labeling was not sufficient for our purpose. Our hypothesis was that the PEG spacer amino groups failed to react with FITC because of interference from the nearby DTPA groups. These DTPA groups may enhance the degree of protonation of primary amine groups on the spacer by creating a strongly anionic environment that can interact electrostatically with the protonated amine. For example, in a previous, unrelated experiment, a DTPA polymer was incubated in an aqueous buffer of 4-methoxybenzylamine/4-methoxybenzylammonium chloride. Upon purifying the polymer in a spin filter with water washes, we found that the 1H NMR spectrum showed 1.7 equivalents of 4-methoxybenzylammonium per DTPA unit. On further washing in a spin filter with phosphate buffer, we found that the 4-methoxybenzylammonium signals had disappeared from the 1H NMR spectrum. The organic amine had been replaced by sodium counterions. This evidence shows that the DTPA polymer electrostatically retains cations despite water washes in a spin filter. Similarly, Plamper and coworkers found that specific cationic counterions are not removed from poly(acrylic acid) by dialysis against water.20 The polymers reported in this work were purified with phosphate buffer washes. However, the fact that the PEGspacer amino groups are chemically attached to the polymer likely allowed the DTPA groups to retain these amino groups as an ion pair. We explored several options to promote the nucleophilicity of the amines at the end of the PEG spacer groups. Some of our initial attempts at optimization are discussed in the Supporting Information. The optimal approach was to load the polymer with an excess of La3+ ions, with the idea that these ions could be removed by ion exchange at a later stage of the reaction. Following a protocol described previously,12 we utilized loading conditions (1.5 equiv of LaCl3) designed to add more than 1 La3+ ion per DTPA.21 In previous experiments, we observed that a polymer with a DTPA group on every repeat unit precipitated under these conditions.12 In contrast, in the experiments described here, we observed that the DTPA polymer with 12% PEG amino spacer groups remained watersoluble under these conditions. However, the polymer did show an additional higher molecular weight shoulder in the aqueous SEC chromatograph. Also, the main polymer peak shifted to a relatively lower apparent molecular weight; this particular effect has been observed in previous lanthanide-loading reactions.12 Aqueous SEC chromatographs for these two polymers are presented in Figure S4. On reacting this La3+-loaded polymer with 30 equiv of FITC (per polymer chain) in phosphate buffer (200 mM, pH 8.00) with 1 M KNO3 and DMSO (40% v/v), we found 37% dye functionalization (3.4 dyes per chain). Multiple spin filter washes with a sodium DTPA (100 mM, pH 8.0) buffer stripped E

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

tested in three model cytometric bioassays. The goal of the first bioassay was to test the basic mass cytometric performance of the four polymer samples. Each of the four samples, as well as a positive control of commercially available X8 metal-chelating polymer, was separately loaded with a different lanthanide ion and then used to label a secondary antibody directed against a murine monoclonal IgG specific for CD34. To start, an aliquot of secondary goat antimouse (GAM) was treated with TCEP to partially reduce disulfides in the hinge region of the antibody. The resultant thiol groups were reacted to form covalent bonds with the maleimide group of the given metal-chelating polymer. The five GAM polymer-antibody conjugates were set aside. Samples of growing KG1a and EL4 cells were collected from their tissue culture flask and stained with CD34 primary antibody, after which one of the GAM-tags was used to stain the Fc fragment of CD34. Prior to mass cytometric analysis, the cells were fixed and incubated with an iridium-containing DNA intercalator in order to identify cell events.15 Representative results for the mass cytometric analysis of the stained KG1a cells are presented in Figure 2. In parts (A) and

groups of the polymer as well as the particular maleimide used to capture the thiol on the polymer chain end. While we do not at the present time understand why the maleimide yields for the DyLight polymers described in this paper are lower than those of the dye-free DTPA polymer, they are sufficient for us to proceed with experiments to test the performance of these polymers as tagging reagents for affinity bioassays. Fluorescence Spectra. As a final step for use in mass cytometric bioassays, the polymers were loaded with ytterbium isotopes and analyzed by fluorescence spectroscopy. Ytterbium was chosen because it has no visible absorbance or fluorescence, although other lanthanides are not likely to be problematic. The metal-loaded maleimide polymers were analyzed by fluorescence spectroscopy to confirm that attachment of the dyes to the polymer did not have any adverse or unexpected effects on the excitation and fluorescence spectra of the dyes. Both excitation and emission spectra conform to literature (FITC)16 or manufacturer specifications (DyLight), as presented in Figure 1. Mass Cytometry and FACS Affinity Bioassays. Proof of Concept Bioassays. To begin, the dual-purpose polymers were

Figure 2. Mass cytometry results for the basic test of mass cytometry performance of the four dual-purpose polymers. KG1a cells were first stained with CD34 and, after washing, were subsequently stained with a secondary goat antimouse labeled with metal-chelating polymer at a concentration of 10 μg/mL. (A, C) 191Ir vs 193Ir is plotted to select for cell events, for the DyL549-Ho165 and the X8-Tb159 tags, respectively. (B, D) Histogram of lanthanide signal is plotted for the cell events from (A) and (C), respectively. Both cells show positive response, although the X8-tagged cell has a higher response. Notably, the DyL549 trial shows a fraction of cells with a low level of lanthanide loading. Figure 1. Normalized fluorescence spectra of P(12%PEGAmino)(88% DTPA)(Yb)(DYE)-maleimide polymer samples. Fluorescence spectra were collected in phosphate buffer (200 mM, pH 8.00). (Top) Fluorescence excitation spectra. Emission was monitored at wavelengths of 450/550/600/690 nm for DyL450/FITC/DyL549/ DyL649, respectively. (Bottom) Fluorescence emission spectra. The dyes were excited at wavelengths of 370/450/510/620 nm for DyL450/FITC/DyL549/DyL649, respectively.

(C), cell events were selected for the DyL549-Ho165 and X8Tb159 trials, respectively. In parts (B) and (D), a histogram of metal intensity was plotted for the previously selected cell gate. Note that the histogram for the X8 polymer in (D) is quite clean, with nearly zero cells containing low lanthanide signal. Conversely, the histogram for the DyL549 polymer shows a fraction of cells with low lanthanide signal. This histogram is F

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

Figure 3. Summary of mass cytometry results for the basic test of mass cytometry performance of the four dual-purpose polymers. KG1a cells are positive for CD34 and thus serve as the positive control, whereas EL4 cells are negative for CD34 and thus serve as the negative control. All four fluorescent polymer-antibody conjugates show moderate metal intensity, although not as high as that obtained with the control X8 polymer-antibody conjugate. The FITC and DyL405 conjugates yield undesired high background on the EL4 cells.

representative of results with the other three fluorescent GAMtags as well. Data for all four fluorescent GAM-tags is summarized in Figure 3. For the positive KG1a cells the four fluorescent conjugates showed moderate metal intensity, although not as high as that obtained with the control X8 conjugate. Note that it is not entirely accurate to compare relative metal intensities between different lanthanide isotopes. This inaccuracy arises from the relationship between metal intensity and the actual number of atoms per cell. The number of metal atoms per cell is calculated from the metal intensity through multiplying by an intensity-to-count conversion factor and dividing by a transmission coefficient.15 These instrument corrections are different for different isotopes, vary between instruments, and will vary for the same instrument over time. Thus, while the comparison to the X8 conjugate is not strictly quantitative, the large difference in response nevertheless demonstrates that the fluorescent conjugates are less efficient at labeling the cells. The other salient result from Figure 3 is that the FITC and DyL405 conjugates showed an undesirable high background with the negative EL4 cells. These results indicated that the fluorescent tags are not as sensitive as the control X8 tags and, in some cases, display undesirable nonspecific adsorption to cells. Nevertheless, we felt the initial results were encouraging enough to justify further testing in functional assays. Our second and third proof of concept assays involved testing the FITC polymer in both mass cytometric and FACS assays. P(12%PEGAmino)(88%DTPA)(172Yb)(FITC)-maleimide (FITC-Yb172) was used to label another aliquot of goat antimouse (GAM), as described above. This antibody GAM-tag was set aside. Growing Ramos cells were collected from the tissue culture flask and stained with CD45 primary antibody, after which the GAM-tag was used to stain the Fc fragment of CD45. As above, prior to mass cytometric analysis, the cells were fixed and incubated with an iridium-containing DNA intercalator in order to identify cell events. Results for the

mass cytometric analysis of the stained Ramos cells are presented in Figure 4A,B. Satisfactory 172Yb signal was obtained when the cells were stained with a 2.5 μg/mL solution of the GAM-tag. Unlike the previous trial, we did not observe a fraction of cells in the histogram with low lanthanide signal. In a separate experiment, a fresh batch of cells was stained with CD45 and the same 2.5 μg/mL solution of GAM-tag, and then analyzed by FACS on a BD Biosciences LSRII instrument. Unstained Ramos cells were also run on the instrument as a negative control. Results from these assays are presented in Figure 4C,D. In Figure 4C, side-scatter is plotted against forward-scatter to gate for intact cells in the stained sample. In Figure 4D, a histogram of the fluorescent signal due to fluorescein is plotted for the unstained and stained lymphocyte populations. The stained cells show a positive response well above the level of signal from the unstained cells. These results show that the GAM-tag prepared with P(12% PEGAmino)(88%DTPA)-(172Yb)(FITC)-maleimide generated both elemental and fluorescent signals for mass and flow cytometry, respectively. Furthermore, cells stained with the GAM-tag were also identified by these readouts. Therefore, our next step was to employ all four polymers in primary antibody tetraplex assays. Preparation of Primary Antibody Tags. The performance of all four dual-purpose polymers was tested in primary antibody assays of KG1a and Jurkat cells. Four different metallabeled primary antibodies were created using the same TCEP strategy described above. Table 2 lists the tag names, primary antibody, and respective polymer tag and metal isotope. In columns 5 and 6, the table also includes values published by Ornatsky and co-workers15 in 2010 for the relative metal intensities of the four antigens for each of the two cell lines. These data were determined by a multiplexed mass cytometric analysis of KG1a and Jurkat cells, using metal-chelating polymer that was commercially available from DVS Sciences. In the antibody labeling step, the antibodies after reduction of the disulfide in the hinge region were treated with an excess G

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

labeling step, we estimate the number of metal atoms per antibody. If the number of metal atoms per polymer is determined independently,2 one can calculate the mean number of polymer molecules attached to each antibody. In the protocol used in previous experiments,2,15 the concentration of metal ions in each antibody solution was determined by ICP-MS, and the concentration of antibodies was estimated from the UV absorbance of the solution at 280 nm, as determined on a Nanodrop instrument. While the ICPMS analysis worked well for the antibody solutions examined here, the presence of the dyes on the polymers attached to the antibodies contributed to the absorbance at 280 nm. This extra absorbance made the Nanodrop-determined antibody concentrations inaccurate; thus, it was necessary to find an alternative method of quantifying antibody concentration. Instead of a direct quantification, we chose to use antibody concentration data from separate antibody labeling reactions. On the same day that we prepared the four fluorescent tags, we used the same protocol to also prepare eight other labeled antibodies with (nonfluorescent) metal-chelating polymers. Of the eight antibodies, three were with CD3, two were with CD34, one was with CD38, and two were with CD45. For our calculations with the fluorescent CD3, CD38, and CD45 tags, we used the respective average antibody concentrations of the CD3, CD38, and CD45 tags prepared with nonfluorescent reagents. For our calculation with the fluorescent CD13 tag, we used the average antibody concentration of all eight antibodies prepared with the nonfluorescent reagents. The results with this approach are presented in column 4 of Table 2. The values range from 135 to 271 metal atoms per antibody. These values are reasonable, and compare reasonably well with the result of 161 atoms per antibody for the GAM-tag described previously.2 Because these values were determined without direct determination of the antibody concentrations, these dye-labeled antibody−polymer conjugates cannot be used in quantitative mass cytometric bioassays.2 Mass Cytometry Antibody Dilution Series and Tetraplex Assay. Mass cytometry experiments began with a tetraplex antibody dilution series. Six titration solutions were prepared, where each staining solution contained a different concentration of all four metal-labeled antibodies. Concentrations of each primary antibody tag in each staining solution are listed in Table S1. In the assay, separate aliquots of KG1a and Jurkat cells were stained with each staining solution and subsequently analyzed by mass cytometry. From the results, one can

Figure 4. Mass cytometry and FACS results for proof of concept bioassays using the secondary antibody GAM conjugate of FITCYb172 to stain Ramos cells previously stained with CD45. Mass cytometry: (A) 191Ir vs 193Ir is plotted to select for cell events. This is shown for the experiment with GAM-tag at a concentration of 2.5 μg/ mL. (B) A histogram of 172Yb signal is plotted for the cell events from (A). High signal was obtained when the cells were stained with a 2.5 μg/mL solution of GAM-tag. FACS: (C) Side-scatter vs forwardscatter is plotted to gate for intact cells in the sample. (D) A histogram of fluorescent signal due to fluorescein is plotted for the previously gated lymphocyte populations. The unstained cells show a level of signal comparable to background, whereas the stained cells show positive response.

of polymer with a terminal maleimide group. Excess polymer was removed by careful washing using spin filters with a 50 kDa cutoff. Due to the low maleimide functionality of the polymers described in this paper, we were initially concerned that there might be insufficient labeling of the primary antibodies with polymer chains. This is a problem that can in principle be overcome by using a larger excess of polymer in the antibody labeling step. To monitor the effectiveness of the polymer

Table 2. Primary Antibodies, Respective Polymer and Metal Isotopes, Metal Atoms per Antibody, and Expected Relative Metal Intensities for KG1a and Jurkat Cells of the Four Primary Antibody Tags tag name CD3-Sm152FITC CD13-Tm169DyL405 CD38-Ho165DyL549 CD45-Tb159DyL649

primary antibody CD3 CD13 CD38 CD45

metal atoms per antibodya

polymer and metal isotope P(12%PEGAmino)(88%DTPA)(152Sm) (FITC)-maleimide P(12%PEGAmino)(88%DTPA)(169Tm) (DyL405)-maleimide P(12%PEGAmino)(88%DTPA)(165Ho) (DyL549)-maleimide P(12%PEGAmino)(88%DTPA)(159Tb) (DyL649)-maleimide

KG1a expected relative metal intensitiesb

Jurkat expected relative metal intensitiesb

135

4

211

182

97

4

271

190

323

159

1000

1000

a Determined by solution ICP-MS analysis to determine metal content and Nanodrop analysis of similar nonfluorescent antibody tags to provide an estimate of antibody concentration. bRelative metal intensities of each antigen for the given cell line. This data was determined by a multiplexed mass cytometric analysis using commercially available metal-chelating polymer from DVS Sciences, performed by Ornatsky and co-workers in 2010.15 In their analysis, CD3, CD13, CD38, and CD45 primary antibodies were labeled with polymers carrying 152Sm, 166Er, 165Ho, and 159Tb, respectively.

H

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

Figure 5. Comparison of mass cytometry relative metal intensities for KG1a and Jurkat Cells to published data.15 Relative intensities were calculated by dividing the metal counts obtained for each tag with the highest concentration antibody staining solution by the metal counts for the CD45-tag, and multiplying by 1000. The relative metal intensities are plotted next to data for relative metal intensities reported in ref 15.

though the other three tags were labeled with the same metal isotope, there is also the concern that the instrument corrections will vary over time. Thus, the comparison of relative metal intensities presented in Figure 5 should be interpreted only in a semiquantitative manner. The cell-labeling efficiency of each conjugated antibody can be estimated from the data in Figure 5. Consider in detail the results for the CD3-tag. The relative metal intensities obtained for KG1a cells in this work and in the work by Ornatsky et al. are both near zero. Therefore, we conclude that the near zero signal obtained in this work is due to lack of CD3 antigen expression on KG1a cells. The relative metal intensity obtained for Jurkat cells in this work closely matches that found by Ornatsky et al. From this, we conclude that the CD3-tag described here efficiently labeled Jurkat cells. Using similar analyses, we reached conclusions about the other antibody tags. The CD13-tag described in this paper seems to overlabel both the KG1a and Jurkat cells. This suggests a problem with this tag; the tag might be undergoing nonspecific adsorption with the cells. This is further evidence of the nonspecific adsorption seen with the DyL405 polymer in Figure 3. Inversely, the CD38-tag described in this paper shows much lower than expected signal. Therefore, we conclude that the CD38-tag did not efficiently label cells. In total, these mass cytometry experiments demonstrated that the CD3 and CD45 tags were efficient at labeling cells, while the other two tags were not. The next step in the evaluation of the antibody tags was a FACS assay. FACS Tetraplex Assay. A tetraplex FACS assay was performed identically to the mass cytometry assay described above. Fresh cells from the same batches of KG1a and Jurkat cells were stained with the highest concentration antibody staining solution, and then run on the FACS instrument. Unstained KG1a cells were also run as a negative control. Data from this assay is presented as histogram plots in Figure 6. We do not show the side-scatter versus forward-scatter plot used to select for cells, as it is similar to that shown in Figure

determine what antibody concentration, if any, yields maximum intensity. Results for the dilution series are presented in Figure S10 in the Supporting Information. In the figure, one can see that the level of metal intensity leveled off as the concentration of labeled antibody became sufficiently high and the cell reached its antibody-binding capacity.26 Both cells showed a high response for the CD45-tag, a moderate positive response for the CD13-tag, and only the Jurkat cells showed a moderate positive response with the CD3-tag. Neither cell showed a positive response with the CD38-tag. Generally, metal intensity of at least 1000 counts per cell is expected for a highly expressed antigen such as the CD45 antigen. The high response of the CD45-tag demonstrates that this labeled antibody functioned well in this bioassay. On the other hand, the metal intensities corresponding to the other three tags were significantly lower than 1000 counts per cell. It is not clear whether these lower counts are due to lower relative antigen expression on the cells15 or due to inefficient labeling of the cell by the antibody. To separate these two factors, we present the data in a different format in Figure 5. Figure 5 contains selected data from Figure S10 in the form of relative metal intensities. This figure was prepared as follows. First, we began with the metal intensities from Figure S10 that were obtained with the highest concentration of each antibody tag. Next, we divided the metal intensities for each tag by the metal intensity for the CD45-tag, and multiplied by 1000. (In other words, the metal intensities for the CD45-tag were used as a normalization factor.) This calculation yielded the relative intensities for each labeled antibody. Finally, the relative intensities for each labeled antibody were plotted next to the relative metal intensities calculated from the data of Ornatsky et al.15 As mentioned in the discussion of the results in Figure 3, it is difficult to directly compare metal intensities for different isotopes, and for data collected at different times. This is a small concern for the CD13-tag because it was labeled with 166Er in the work by Ornatsky et al., but 169Tm in this work. Even I

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

Finally, the CD45 response is plotted in Figure 6D. Both cell lines show fluorescence intensity significantly higher than that of the unstained negative control. The high fluorescence intensity of the CD45+ cells is further evidence that the CD45− tag efficiently labeled cells. In total, these results from the FACS assay add further support to the conclusions reached from the mass cytometry experiments. Namely, the CD3 and CD45 tags were efficient at labeling cells, while the CD13 and CD38 tags were not. Poor Tag Performance. In contrast to the work reported here, previous experiments with a number of antibodies, including CD3, CD38, and CD45, labeled with an analogous metal-loaded polymer of identical length carrying only DTPA pendant groups, indicated that these labeled antibodies were effective at antigen recognition in mass cytometry experiments.2 We suspect that the difficulties we encountered with the DyL405- and DyL549-containing polymers may be caused by the particular dyes themselves. It is known, for example, that when the dye Texas Red is conjugated to antibodies, the dye antibody conjugates are sticky and prone to agglutination.28 On the other hand, if Texas Red is first conjugated to BSA, the Texas Red−BSA conjugate can be attached to an antibody to form a stable adduct with very little nonspecific binding.29 In another example, Bergman and co-workers describe silica particles with a surface polyethyleneimine coating, to which streptavidin labeled with either Alexa 555 or DyLight 549 was covalently bound. They expected that as the pH was raised, the polyethyleneimine groups would become deprotonated (neutral) and consequently the silica particles would agglomerate. As they raised the pH, they found that the DyLight 549 sample agglomerated above pH 6.5, whereas the Alexa 555 sample agglomerated above pH 8.30 The relative instability of the DyLight 549 sample indicates that different dyes can have a profound effect on the colloidal stability or stickiness of a given system. Given these points, we believe that further experiments should focus on other reactive fluorescent dyes.

Figure 6. FACS tetraplex assay with four dual-purpose antibody tags and KG1a and Jurkat cells. (A−D) Histograms of (fluorescent) FACS signal are plotted for the cell events, after gating for cell events as described in Figure 4C. The CD13 and CD38 tags fail to show any signal above a background level, the CD3 tag identifies CD3+ and CD3− populations in the Jurkat cells, and the CD45 tag shows positive signal for both cell samples.

4C. In Figure 6A−D, histogram plots of fluorescence intensity are presented for each antibody tag. The x-axis denotes the fluorescence intensity for each cell event, and the y-axis denotes the relative number of cell events with a given fluorescence intensity. These figures are interpreted by comparing the fluorescence intensity of the stained cells to that of the unstained KG1a. The unstained KG1a was included as a negative control to account for cell autofluorescence. Distinct peaks are termed cell populations, and cell populations with high and low levels of signal are considered + and − for the given antigen, respectively. The CD3 response is plotted in Figure 6A. All the KG1a cells had near the same fluorescence intensity as the unstained negative control. The majority of the Jurkat cells had the same level of intensity; however, there was a small population with a higher level of fluorescence. From this result we concluded that KG1a cells were CD3-, and Jurkat cells were a mix of CD3+ and CD3−. One might ask why only 10−20% of the Jurkat cells appear to be CD3+. When a similar histogram is constructed for the mass cytometry data (not shown), it appears that approximately 40% of the cells are CD3+. In the literature, there is a report27 of a sample of Jurkat cells in which 57% of the cells are CD3+. It is possible that the discrepancy between the two assays described here is simply due to each assay being run with Jurkat cells grown to different densities. Figure 6B,C contain the data for the CD13 and CD38 response. In both plots, the stained cells show near the same level of fluorescence as the unstained negative control. This lack of positive response stands as further evidence that the CD13 and CD38 tags did not perform efficiently.

■

CONCLUSIONS Four dual-purpose metal-chelating polymers were synthesized from a common precursor polymer containing on average nine amino PEG spacers and seventy DTPA groups per chain. The reaction for the attachment of fluorescent dyes was optimized to yield polymers with 2.6 to 6.2 dyes per chain. The dyes employed in this study were FITC and DyLight dyes 405/549/ 649. In the final step of the synthesis, each dye-labeled polymer was end-functionalized with a bismaleimide linker. Unfortunately, the degree of maleimide functionalization was lower than that observed for similar, nonfluorescent polymer samples. An initial mass cytometric bioassay was performed with all four fluorescent polymers, as well as a commercially available control polymer (X8). All five polymers were covalently attached to GAM. The GAM-tags were used to stain KG1a (positive) and EL4 (negative) cells previously stained with CD34, which were analyzed by mass cytometry. The fluorescent tags did impart lanthanide signal to the KG1a cells, but showed lower signal than the X8-tag. Additionally, two of the fluorescent tags displayed nonspecific adsorption to the EL4 cells. To test the feasibility of a dual-purpose tag, the FITCcontaining polymer was covalently attached to GAM. The GAM-tag was used to stain Ramos cells previously stained with CD45, which were analyzed both by mass cytometry and J

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

■

FACS. Happily, the GAM reagent showed positive signal by both methods. The four polymer samples were used to create primary antibody-tags with CD3, CD13, CD38, and CD45. These antibodies were chosen as specific analytes for two cell lines of interest, KG1a and Jurkat. Aliquots of both cell lines were stained with all four antibodies and analyzed by mass cytometry. The results indicated that the CD3 and CD45 tags were efficient at labeling cells, but the CD38 tag was not, and the CD13 showed signs of nonspecific adsorption. Next, aliquots of both cell lines were stained with all four antibodies, and then analyzed by FACS. The results further supported the conclusions from the mass cytometry experiments. These results are bittersweet. The positive results with the CD3 and CD45 tags demonstrate that a dual-purpose polymer can be used to create an antibody tag that carries both elemental and fluorescent markers. On the other hand, the negative results with CD13 and CD38 tags point to problems with the DyL405and DyL549-containing polymers used to create the antibody tags. Results from both the initial proof of concept assays as well as from the primary antibody assays all point to the borderline performance of these polymers. Further experiments with other reactive fluorescent dyes (for example, Alexa 555) are necessary to understand whether the dyes themselves interfere in the assays, or whether the synthetic polymer strategy must be revisited. Future work should also be directed toward optimization of the maleimide functionality of the polymer. In this regard, we note that polymers bearing a terminal furan group will also react with excess bismaleimide to install a terminal maleimide group.31

■

REFERENCES

(1) Bandura, D. R.; Baranov, V. I.; Ornatsky, O. I.; Antonov, A.; Kinach, R.; Lou, X.; Pavlov, S.; Vorobiev, S.; Dick, J. E.; Tanner, S. D. Anal. Chem. 2009, 81, 6813−6822. (2) Majonis, D.; Herrera, I.; Ornatsky, O.; Schulze, M.; Lou, X.; Soleimani, M.; Nitz, M.; Winnik, M. A. Anal. Chem. 2010, 82, 8961− 8969. (3) Bendall, S. C.; Simonds, E. F.; Qiu, P.; Amir, E. -a. D.; Krutzik, P. O.; Finck, R.; Bruggner, R. V.; Melamed, R.; Trejo, A.; Ornatsky, O. I.; Balderas, R. S.; Plevritis, S. K.; Sachs, K.; Pe’er, D.; Tanner, S. D.; Nolan, G. P. Science 2011, 332, 687−696. (4) Shapiro, H. M. Practical Flow Cytometry; Alan R. Liss: New York, 1985. (5) Alexander, C. M.; Puchalski, J.; Klos, K. S.; Badders, N.; Ailles, L.; Kim, C. F.; Dirks, P.; Smalley, M. J. Cell Stem Cell 2009, 5, 579−583. (6) Osawa, M.; Hanada, K.; Hamada, H.; Nakauchi, H. Science 1996, 273, 242−245. (7) Van Bekkum, D. W.; Van den Engh, G. J.; Wagemaker, G.; Bol, S. J.; Visser, J. W. Blood Cells 1979, 5, 143−159. (8) Zhang, Z.; Yan, X.; Xu, M.; Yang, L.; Wang, Q. J. Anal. At. Spectrom. 2011, 26, 1175−1177. (9) Purvis, N.; Ornatsky, O.; Shults, K.; Baranov, V.; Tanner, S.; Stelzer, G.A Comparative Analysis of Quantitative Techniques using Flow and Mass Cytometric Platforms. Presented at The XXV Congress of the International Society for Advancement of Cytometry, Seattle, WA, USA, May 8−12, 2010, ISAC: Bethesda, MD, 2010; p P289. (10) Paik, C. H.; Murphy, P. R.; Eckelman, W. C.; Volkert, W. A.; Reba, R. C. J. Nucl. Med. 1983, 24, 932−936. (11) Louie, A. Chem. Rev. 2010, 110, 3146−3195. (12) Majonis, D.; Ornatsky, O.; Kinach, R.; Winnik, M. A. Biomacromolecules 2011, 12, 3997−4010. (13) Gottlieb, H. E.; Kotlyar, V.; Nudelman, A. J. Org. Chem. 1997, 62, 7512−7515. (14) Lou, X.; Zhang, G.; Herrera, I.; Kinach, R.; Ornatsky, O.; Baranov, V.; Nitz, M.; Winnik, M. A. Angew. Chem., Int. Ed. 2007, 46, 6111−6114. (15) Ornatsky, O.; Bandura, D.; Baranov, V.; Nitz, M.; Winnik, M. A.; Tanner, S. J. Immunol. Methods 2010, 361, 1−20. (16) Klonis, N.; Sawyer, W. H. J. Fluoresc. 1996, 6, 147−157. (17) Agi, Y.; Walt, D. R. J. Polym. Sci., Part A: Polym. Chem. 1997, 35, 2105−2110. (18) Manning, T. J.; Fiskus, W.; Mitchell, M.; Dees, L. Spectrosc. Lett. 1999, 32, 463−467. (19) Lavis, L. D.; Rutkoski, T. J.; Raines, R. T. Anal. Chem. 2007, 79, 6775−6782. (20) Plamper, F. A.; Becker, H.; Lanzendörfer, M.; Patel, M.; Wittemann, A.; Ballauff, M.; Müller, A. H. E. Macromol. Chem. Phys. 2005, 206, 1813−1825. (21) Ternovaya, T. V.; Kostromina, N. A.; Kundrya, V. T. Zh. Neorg. Khim. 1975, 20, 2957−63. (22) Thermo Scientific. Prod. Nos. 46426, 46400, 46402, 46407, 46412, 46414, 46415, 46418, 46420, 46421 53068 Doc. No. 1963 Instructions for DyLight Amine Reactive Fluors. (23) Friedman, M.; Wall, J. S. J. Org. Chem. 1966, 31, 2888−2894. (24) Epps, D. Anal. Biochem. 2001, 295, 101−106. (25) Miyadera, T.; Kosower, E. M. J. Med. Chem. 1972, 15, 534−537. (26) Davis, K. A.; Abrams, B.; Iyer, S. B.; Hoffman, R. A.; Bishop, J. E. Cytometry 1998, 33, 197−205. (27) Zeng, H.; Wang, H.; Chen, F.; Xin, H.; Wang, G.; Xiao, L.; Song, K.; Wu, D.; He, Q.; Shen, G. Anal. Biochem. 2006, 351, 69−76. (28) Tsurui, H.; Nishimura, H.; Hattori, S.; Hirose, S.; Okumura, K.; Shirai, T. J. Histochem. Cytochem. 2000, 48, 653−662. (29) Roederer, M. Texas Red-BSA Conjugation of Antibodies, http://www.drmr.com/abcon/trbsa.html (accessed February 2013). (30) Bergman, L.; Rosenholm, J.; Ö st, A.-B.; Duchanoy, A.; Kankaanpäa,̈ P.; Heino, J.; Lindén, M. J. Nanomater. 2008, 2008, 1−9. (31) Illy, N.; Majonis, D.; Herrera, I.; Ornatsky, O.; Winnik, M. A. Biomacromolecules 2012, 13, 2359−2369.

ASSOCIATED CONTENT

S Supporting Information *

Experimental details for the polymer synthesis, details on the initial attempts to optimize the dye-labeling reaction, an expanded discussion on attachment of the bismaleimide linker, table of antibody tag concentrations in the staining solutions used for the tetraplex antibody dilution series, 1H NMR and gCOSY spectra of P(12%PEGAminoBoc)-disulfide (D2O), 1H NMR spectrum of P(12%PEGAmino)(88%DTPA)-disulfide (D2O), aqueous SEC chromatographs of P(12%PEGAmino)(88%DTPA)-disulfide, P(12%PEGAmino)(88%DTPA)-disulfide-La3+ loaded, and P(12%PEGAmino)(88%DTPA)(FITC)disulfide, normalized UV/vis absorption spectra of P(12% PEGAmino)(88%DTPA)(Dye)-disulfide polymers, partial 1H NMR spectra of P(12%PEGAmino)(88%DTPA)(Dye)-maleimide (D2O) polymers, and mass cytometry antibody dilution series with the four dual-purpose antibody tags. This material is available free of charge via the Internet at http://pubs.acs.org.

■

Article

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Fax: 1-(416) 978-0541. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank NSERC Canada, the Province of Ontario, and DVS Sciences for their support of this work. D.W. thanks the Alexander von-Humboldt foundation for a Feodor-Lynen fellowship. K

dx.doi.org/10.1021/bm4001662 | Biomacromolecules XXXX, XXX, XXX−XXX