Liquid Chromatography Separation and Mass Spectrometry Detection

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Liquid Chromatography Separation and Mass Spectrometry Detection of Silver-Lipoate Ag29(LA)12 Nanoclusters: Evidence of Isomerism in the Solution Phase David M. Black, Geronimo Robles, Priscilla Lopez, Stephan B. H. Bach, Marcos M. Alvarez, and Robert L. Whetten Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b04104 • Publication Date (Web): 20 Dec 2017 Downloaded from http://pubs.acs.org on December 21, 2017

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Analytical Chemistry

Liquid Chromatography Separation and Mass Spectrometry Detection of Silver-Lipoate Ag29(LA)12 Nanoclusters: Evidence of Isomerism in the Solution Phase David M Black a*, Geronimo Robles a, Priscilla Lopez a, Stephan B H Bach b, Marcos Alvarez a, Robert L Whettena* Departments of aPhysics & bChemistry, University of Texas, San Antonio, TX 78249, USA

ABSTRACT

Evidence for the existence of condensed phase isomers of silver-lipoate clusters, Ag29(LA)12, where LA = (R) - α lipoic acid, was obtained by reversed-phase ion-pair liquid chromatography with inline UV-vis and ESI-MS detection. All components of a raw mixture were separated according to surface chemistry and increasing size via reversed-phase gradient HPLC methods and identified by their corresponding m/z ratio by electrospray ionization (ESI) in the negative ionization mode.

Aqueous and methanol mobile-phase mixtures, each containing 400 mM

hexafluoroisopropanol (HFIP) – 15 mM triethylamine (TEA), were employed to facilitate

ACS Paragon Plus Environment

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interaction between the clusters and stationary phase via formation of ion-pairs. TEA-HFIP (triethylammonium-hexafluoroisopropoxide)

had

been

shown

to

provide

superior

chromatographic peak shape and mass spectral signal compared with alternative modifiers such as TEAA (triethylammonium-acetate) for analysis of oligonucleotide samples.

Liquid

chromatographic separation prior to mass spectrometry detection facilitated sample analysis by production of simplified mass spectra for each eluting cluster species and provided insight to the existence of at least two major solution-phase isomers of Ag29(LA)12. UV-Vis detection in-line with ESI analysis provided independent confirmation of the existence of the isomers and their similar electronic structure as judged from their identical optical spectra in the 300-500 nm range. [Diastereomerism provides a possible interpretation for the near-equal abundance of the two forms, based on a structurally defined non-aqueous homolog.]

1. INTRODUCTION Noble metal monolayer-protected clusters (MPCs) have the potential to play significant roles as novel molecules for a range of applications. MPCs of varying size and stoichiometry exhibit interesting optical,1,2 electrochemical,3-5 catalytic,6,7 magnetic8,9 properties as well as tunable solution-phase properties that can be customized via the incorporation of organic ligands10,11 that impart pre-engineered solubility, stability, acid-base character, salt form, and functional capability (i.e., biological activity).12,13 Larger silver nanoparticles, are known to exhibit widespectrum antibiotic activity against bacteria – and biofilms – that have acquired resistance to conventional antibiotics.14-19

The ability to design and synthesize nanoclusters, of definite

stoichiometry, that combine a silver core with ligands of demonstrated antibiotic activity, will make it possible to investigate a range of MPC compositions for use as novel therapeutics.


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Traditionally, noble-metal cluster characterization has been carried out by a combination of tools including microscopy,20 crystallography, spectroscopy (NMR, UV-Vis, IR, Raman), analytical ultracentrifugation, dynamic light scattering/zeta-potential,21 and direct infusion mass spectrometry.22,23 These methods provide measures of the metal core, the cluster – to – ligand stoichiometry, ligand structure and cluster mass. Methods such as these perform best when sample purification and pre-concentration is carried out prior to analysis. For that purpose, polyacrylamide gel electrophoresis (PAGE) is routinely used for separation of MPC product mixtures prior to analysis.24 However, PAGE suffers from relatively low separation efficiencies, is not particularly effective for hydrophobic compounds, and requires sample work-up to remove separated components from the gel after separation (for subsequent investigation by other techniques), which may be relatively labor intensive. Separation of MPC synthetic products by size exclusion25,26, thin layer27 and reversed-phase28 chromatography methods have been reported as an effective alternative to gel electrophoresis for analysis of these complex molecules. Early reversed-phase chromatography was reported by Murray and coworkers,28-31 who used C8 stationary phases, coupled with photodiode array and electrochemical detection, to successfully resolve non-aqueous Au-MPCs.

More recently,

Negishi and coworkers advanced this work using preparatory and conventional flow rate reversed-phase LC combined with UV detection to demonstrate the separation of gold and alloy MPCs over a wide range of core diameter32 and charge,33 ligand structures,34 and coordination isomer35. Bürgi and coworkers were successful at separation of the chiral Au38(SR)24 via use of chiral LC columns.36 In all these instances, fraction collection of the LC eluent permits each of the separated fractions to be identified compositionally by subsequent MS analysis. Recent work in our lab has shown the advantages of capillary flow in-line liquid chromatography mass


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spectrometry, which negates the need to collect fractions, for analysis of organo-soluble and aqueous clusters.37-39 In 2015, AbdulHalim et al. reported a major advance in the understanding of the silverdithiolate clusters, Ag29(dithiolate)12: They obtained the crystal structure of the non-aqueous compound Ag29(BDT)12(TPP)4 (where BDT = 1, 3 benzene dithiol; TPP = triphenylphosphine) as having four tetrahedrally equivalent positions.40 A thirteen-atom (Ag13) quasi-icosahedral core is encapsulated within an exterior shell composed of four alternating chair Ag3S3 motifs and four Ag1S3P1 crowning motifs. Very recent characterization of this organosoluble cluster by Baksi et al. revealed the existence of isomers in the gas phase of an ion-mobility analyzer.41,42 Parallel to those advances, several groups have reported on the properties of the water-soluble analog stabilized with a di-thiolate lipoate ligand42-45 [ref. 42 concerns BDT, not LA] tentatively identified as Ag29LA12 by analogy to the hydrophobic homolog.40 Although the crystal structure of the water-soluble homolog – the species reported on here – has not yet been reported, a high degree of similarity may be expected between it and the hydrophobic variant for the reason that both incorporate bidentate protecting ligands.

The presence and bonding of a molecule

analogous to the organosoluble counter ion (it's neutral, TPP), TPP, is unknown. Here we demonstrate the use of the inline LC-UV-MS methodology for efficient and sensitive separation and detection of water-soluble Ag29(LA)12, a sample intended for tests of invitro efficacy of its antibiotic and antifungal activity.46 The goal of the work described herein was to develop a rapid (< twenty minute) method for separation and identification of the various components of a sample mixture not subjected to any prior cleanup (i.e., the mother liquor) and, ultimately, to determine relative amounts of each. As the method was optimized, it became evident that the major eluted component resolved into two chromatographic peaks (solution-


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phase populations), each tracking with the intact mass of the Ag29(LA)12 cluster, and subsequently baseline-separated. As the existence of isomers for the non-aqueous cluster was quite recently reported in the gas phase,41 the HPLC results presented here demonstrates the existence of isomers in the solution (liquid) phase. Hence they are not an artifact of the ionmobility method or mass spectrometer analyzer. We show here that analysis via inline LC-MS sample provides information about the composition of the mixture that is not obtainable via direct-infusion mass spectrometry analysis alone. Although this work pertains to acidic watersoluble silver clusters, the technique is also applicable to other cluster varieties (i.e., gold core, organosoluble ligands, etc.).

Additionally, the inline separation makes possible gas-phase

interrogation of these clusters via in-source collisional activation. Ion activation techniques have been reported to provide a means for partial or complete structural elucidation of unknown cluster compositions,22,47 and is used here to support the identification of the two isobaric chromatographic peaks as isomers. 2. EXPERIMENTAL SECTION 2.1 Synthesis and preparation Samples were prepared using a 500 mL-fleaker (Pyrex/Corning) reaction vessel covered with a double layer of aluminum foil. Mixing was carried out using a 2-inch Teflon stirring bar (at low setting). The vessel was capped with aluminum foil after mixing was completed. Ag29LA12 clusters were prepared using a scaled-up version of the preparation described by Linden et al.44 First, the following four solutions, A – D, were prepared using HPLC-grade water for immediate use (not to be stored). A) 1.9 grams of (R)-a-lipoic acid [TCI Cat. L0207 > 98.0 %] were dissolved in 100 mL of HPLC water, adding base to adjust the pH to 8-9 until the acid dissolved. 1 M KOH and 30% NH4OH , and tetraethylammonia (TEA) were used for this purpose. B) 0.70


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grams of NaBH4 was dissolved in 10 mL of HPLC water using a 20 mL capped vial. C) 0.29 grams of AgNO3 was dissolved in 10 mL of HPLC water using a 20 mL vial covered with aluminum foil. D) 1.00 grams of NaBH4 was dissolved in 10 mL of HPLC water using a 20 mL vial. Next, the first sodium borohydride solution (B) was added to the lipoate solution (A) while stirring. This mixture was allowed to react for fifteen minutes with the top covered by aluminum foil. After fifteen minutes elapsed, the silver nitrate solution, (C), was added and allowed to react for fifteen minutes, again with the top covered by aluminum foil. Last, the final sodium borohydride solution, (D), was added to the reaction mixture and allowed to react for at least four hours (top covered with aluminum foil). For the sample prepared here, the reaction was allowed to run overnight under dark conditions. After reaction was allowed to proceed to completion, aliquots of the reaction mixture were diluted – 10x – in mobile phase A (400 mM HFIP – 15 mM TEA) for subsequent LC-MS or LC-UV analysis. 2.2 HPLC-MS and UV-Vis method conditions Liquid chromatography (LC) experiments were performed on an Eksigent nanoLC 2D system coupled to a Bruker micrOTOF time-of-flight mass spectrometer (MS). All separations were carried out using an Ace 300Å C18 HPLC column (0.5 mm x 150 mm, 3 µm particle size) (Advanced Chromatography Technologies Limited, Aberdeen, UK) maintained at ambient laboratory temperature. Mobile phases were prepared 400 mM hexfluoroisopropanol (HFIP) 15 mM triethylamine (TEA) in ddH2O (mobile phase A) and neat methanol (mobile phase B). All solvents for direct infusion and LC-MS were obtained from Fisher Scientific (Fairlawn, NJ). The flow rate used for all experiments was ten microliters per minute (µL/min). Injections – 5.0 µL – were carried out by an Eksigent AS-1 autosampler configured with a 20-µL sample loop. All reaction mixture samples were diluted 20x in mobile phase A. Direct infusion was carried


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out by loop injection (i.e., no column between autosampler and mass spectrometer) using a mobile phase composition of 95% MP A: 5% MP B.HPLC experiments were carried out using twenty-minute linear gradient methods with varied starting and ending mobile phase conditions. After completion of the twenty-minute gradient, 100% methanol was rinsed through the column to remove any non-polar components for five minutes. This was then followed by a twentyminute re-equilibration at initial method conditions. Mass spectrometer acquisition settings were identical for both direct infusion and LC-MS experiments. Data was acquired from m/z 100 6,000. Ten-thousand spectra were summed per spectrum acquired. Nebulizer pressure was set to 4.0 bar. Nitrogen sheath gas was set to zero L/min. The endplate offset and capillary potentials were held at -1000 V and 3500 V, respectively. Capillary exit and skimmer voltage settings were -100 V and -33 V respectively. Lens 1 pre-pulse storage and transfer times were 35 µs and 140 µs, respectively. MCP detector voltage was increased to 2350 V (from 2100 V standard) for improved detection. UV-Vis detection and real-time 240 nm - 700 nm spectral characterization was acquired with an UltiMate™ UV Detector (LC Packings) equipped with a Micro LC (10-100 µL/min) flow cell (UZ-M10, P/N 16011). 3. RESULTS AND DISCUSSION 3.1 Direct Infusion Although direct infusion provides a fast and simple method of sample introduction to the mass spectrometer, there are several factors that may contribute to production of mass spectra that are not representative of the true mixture composition. Peak intensities in mass spectra acquired via electrospray ionization are often poor indicators of the relative abundances of the species present in a solution, (even to the extent that major components of a sample may not be


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evident in a spectrum), and therefore may lead to inaccurate compositional or molecular assignment of the contained species. The reasons for these difficulties may be related to several factors including, varying sensitivity of mass-analyzers at different m/z, differences in ionization efficiency, co-elution of analytes leading to ionization suppression, and adduct formation – and / or fragmentation. Each of these is a common source of added mass spectral complexity which may greatly complicate mass spectral interpretation. Additionally, the presence of isobaric (i.e., isomers) species cannot be detected without separation prior to detection. Any of these factors may affect quantitative or even qualitative interpretation of mass spectra and sample composition. Methods to simplify mass spectra, or to relate signals to one another for purposes of mass spectral interpretation are often essential for correct determination of mixtures. Figure 1 shows a negative ESI mass spectrum obtained from a sample of Ag29(LA)12 analyzed by direct infusion MS. In this spectrum, the doubly-charged parent ion is observed while the triply- and singly-charged ion populations are absent. The observed parent ion charge states – and related fragmentation – may be related to acid and / or base mobile phase modifier selection. Often, ESI fragmentation can be used to good advantage. Here, for instance, we learn that a dithiolate may depart as a neutral disulfide thus preserving charge as the [29,12]2- → [29,11]2peaks in Figure 1 indicate. In this case though, the fragmentation provides few structural details and, in general, the utility of the mass spectrum is restricted by several of the limitations detailed above. Of concern is the poor signal-to-noise ratio which result from a combination of ionization suppression from salt and other small molecule starting materials and reactants, in-source fragmentation that can complicate identification, and solution complexity which is not evident due to the presence of isomers and multimers. Without separation (either by inline LC-MS or by


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fraction collection and subsequent direct infusion), an accurate accounting for all species present is not possible. 3.2 Liquid chromatography-mass spectrometry 3.2.1 Liquid Chromatography The separations discussed here were carried out by capillary-flow (µL/min) ion-pair reversedphase HPLC. Doubly deionized water was selected for mobile phase A and neat methanol for mobile phase B.

These solvents were chosen both for their chromatographic and mass

spectrometric performance. Methanol was found to provide for strong mass spectral signal strength and also for best chromatographic selectivity within the targeted twenty-minute time limit. Separations using acetonitrile in place of methanol were also successful with longer method times and shallower gradients (reference SI section). Alkaline solution conditions, above ~ pH 7.5, are required to achieve a high degree of solubility and efficient ionization of these clusters.

However, highly polar and/or ionized

molecules are not retained by the non-polar aliphatic C18 stationary phase. For this reason, ionpair chromatography was employed to promote interaction between the Ag-MPCs and C18 stationary phase through formation of lipoate – triethylammonium ion-pairs, which have significantly greater affinity for the non-polar alkyl stationary phase (see Scheme 1). The ion paired Ag-MPCs are fully soluble in the mobile phase and exhibit good retention characteristics. The HFIP – TEA combination has been used for analysis of oligonucleotide samples for over two decades and has been reported to produce superior liquid chromatographic and mass spectrometric performance compared with acetic acid and TEA.48-50 Figure 2 shows results from LC-MS separation of 5.0-µL of a ~ 100 ng/µL solution (500-ng total) of the Ag29(LA)12 synthetic mixture in which at four unique components could be


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separated and detected. The figure consists of several overlaid ion traces, each illustrating ion signal intensity as a function of retention time for one of the components in the mixture. The C18 stationary phase provided good separation of the ion-paired clusters while maintaining overall good peak shape, peak width, and signal-to-noise ratio. This trend is consistent with separate reports by Murray and Negishi detailing the use of ion-pair chromatography for separation of water-soluble MPCs.31,51

Hydrophobic interactions between the aliphatic

hydrocarbon portions of the stationary phase and the alkyl portions of the protecting ligands and ion-pairing agent offer an effective mode of separation for the surface chemistry of these clusters. In Figure 2, all major components were retained greater than twice the retention time of the solvent front, and all eluted within twenty minutes. The general trend showed that the compounds eluted in the order of size, beginning with the apparent Ag29(LA)12 isomers at similar retention times: (Isomer #1) at RT 16.3 minutes, followed by Ag29(LA)12 (Isomer #2) at RT 17.0 min, and then by larger dimer- and trimer-like species, ~ Ag61(LA)24 at RT 20.1 min, and RT 21.1 min respectively. Peaks widths, at half height, of the isomer components are approximately 0.2 min. Comparison of the theoretical and experimental isotope patterns for the [Ag29(LA)12]2peaks is shown in Figure S5. Figure 3 shows a comparison of several linear gradient methods varying the starting and organic composition on the effect on RT. Generally, as the organic content of the initial mobile phase conditions in decreased, retention times increased, which is consistent with the mechanism of ion pair reversed-phase LC. Baseline separation of the two isomers is achieved with a shallow gradient, shown in Figure 3b, 30 – 50% MP B over 20 min. Ostensibly, the two Ag29(LA)12 chromatographic peaks correspond to at least two separate solution phase isomers of this cluster species.

Because of the direct relationship between

analyte-to-stationary phase interaction and hydrophobic character of the various separated


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components (also including effects contributed by the triethylamine ion pairing agent), any differences in retention may be attributed to either the number of protecting ligands, or the protecting ligand shell orientation. Here, we observe that one of the Ag29(LA)12 structures is retained more strongly than the other, which indicates slight differences in the interaction / affinity with the C18 stationary phase between the two. We infer from this result that the two structures exhibit different conformations in solution which are likely related to the positioning of one or more of the protecting pMBA ligands. Structural differences between two clusters which result from variances in ligand orientation would be expected to affect how the clusters interacts with the surrounding mobile and stationary phase but would not affect other measurements, including mass and optical absorbance (discussed below).

Reports of MPC

isomerism have been observed by ion mobility for other silver cluster stoichiometries.52 In those cases, ligand orientation and intramolecular hydrogen bonding where postulated as the cause of such isomerization. Those possibilities are equally valid for these apparent solution phase isomers. In Figure S1 is shown an LC-MS chromatogram acquired with the 400 mM HFIP – 15 mM TEA modifiers replaced by 10 mM ammonium acetate (pH 7.5). Although the clusters are ionized and detected by ESI – MS, ammonium acetate is not a strong ion-pairing agent and therefore none of the clusters is retained by the C18 column. 3.2.2 Mass Spectrometry and UV-Vis Detection Matching negative ESI mass spectra for peaks 1 and 2 are shown inset in Figure 2 for comparison purposes. The base peak signals observed in each of the spectra correspond, both by m/z and isotope pattern, to the 2- charge state of Ag29(LA)12. The dimer- and trimer-like components were observed at the 3- to 4- charge states at trace levels. Minimal, if any, TEA or


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HFIP related adducts appear to be present in any mass spectra. Only slight fragmentation was observed for any of these species which made identification of the known components straightforward. Positive mode ESI mass spectra and corresponding tentative peak identifications for the two isobaric species are shown in Figure S4 and Table S1, respectively. Inline UV detection (330 nm) was carried out between the LC and the mass spectrometer. Interestingly, the UV response profile was observed to be qualitatively similar to that produced by electrospray ionization and mass spectrometry detection, as shown in Figure 4.

The

agreement between the two orthogonal detectors may indicate that the relative concentrations of the two isomers is represented in nearly correct proportion by each of these methods and that any differences in ionization efficiency, transmission efficiency and detection efficiency are negligible. This finding may indicate the degree to which these two structures and relative stabilities are similar.

The comparable absorbances indicate near identical electro-optical

structure, which is certainly related to core geometry. Electrospray ionization efficiency and MS signal response is dominated by factors related to solution and gas phase acid / base chemistry. For this reason, we may infer that both isomers (or groups of isomers) have similar surface chemistry (i.e., ligand orientation). It is possible that conformational diastereomers, or some other high-similarity constitutional isomers, would behave in a similar fashion to what is observed here. UV-Vis spectra (240 nm -700 nm) were recorded for each of the two peaks and are shown inset in Figure 4. (The scans were acquired between the leading and tailing edge of each eluting peak at half height and were background subtracted.) The nearly identical spectra serve as practical proof that neither of the two peaks are present because of simple cluster aggregation. Aggregated clusters exhibit structure-less optical spectra and a brown color (instead of the bright


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orange arising from the 425 nm and 500 nm bands). Furthermore, the observation of identical bands (each of which has been modeled to correspond well defined inter-band transitions by Bakr et al.)40 shows that both clusters have similar electronic structure as would be the case if they were structural isomers. The observation of identical optical spectra also minimizes the possibility that the two peaks simply correspond to different oxidation states of the same cluster composition. In that were the case, differing optical spectra would be expected. 3.3 In-Source Fragmentation Representative in-source fragmentation spectra for the two Ag29(LA)12 peaks, acquired via inline LC sample introduction, is shown in Figure 5. Both fragment in nearly identical fashion over a range of capillary exit – to – skimmer voltage differentials (see Figure S3). At relative low (standard) voltage differentials the intact cluster exhibits limited fragmentation such that no more than one dithiooctanoic acid (LA) residue is lost, while maintaining the intact ion population as the majority species in the spectrum. As the voltage differential is incrementally increased, a second dithiooctanoic acid moiety is lost, forming the [29, 10]2- fragment ion structure. Subsequent fragmentation occurs by means of an alternate pathway that forms a cascading series of fragment ions, corresponding to sequential losses of mercaptooctanoic acid neutrals, each maintaining the silver atoms of the parent species. This series extends from the doubly-charged [29, 10]2- fragment ion population to the tentatively identified [29, 3, S7]2fragment ion as shown in Figure 5. The similarity of the fragmentation between each of the isobaric components is suggestive of a high degree of similarity between the gas-phase structures. Table 1 lists the various fragment ions identified in the in-source fragmentation spectrum and corresponding fragmentation processes are illustrated in Scheme 2. Interestingly, all fragment ions exist as even-electron ions.


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4. CONCLUSIONS As shown here, LC-MS offers a sensitive means for fast, high-resolution separation and detection of nanogram-scale quantities of Agn(LA)m clusters ranging 5.6 kDa to 17.5 kDa. The method described here consumes ~ 500 ng of sample per injection for separation and detection of the various mixture components – notably including two, not previously reported, presumed structural isomers which likely arise from differences in conformation of the protecting pMBA ligand shell – with good signal-to-noise ratios. Clusters were separated by ion-pair reversedphase mechanism resulting in the elution of clusters in order of increasing size. Time-of-flight mass spectrometry was used to detect the clusters as they eluted from the HPLC column and to acquire high-resolution mass spectra for identification of each eluting cluster composition. This methodology is applicable to clusters varying in composition, size and surface chemistry – i.e., core metal, and surface ligand chemistry – and is an effective means for determination of synthetic quality via determination of sample composition. The reversed-phase separation of monolayer-protected clusters may also lend itself to various extensions of the chromatographic technique such as quantification of mixture components and measurements of various chemical properties such as pKa, lipophilicity, etc. Future work will be aimed at analysis of mixtures containing clusters of increasing larger core diameters as well as mixtures containing mixed monolayer protected clusters. Further improvements to the chromatography shown here may be obtained through the use smaller particle size and will be investigated in future work.


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O

O OH S

pKa = 4.52

S

O S

-

S

CH3

O

O

+

OH S

S

O

pH = 7.8 TEA/HFIP

S

S

-

HN

CH3

H3C

Scheme 1. Triethylammonium – (R)-α-lipoate ion-pair formation/interaction that solubilizes cluster via deprotonation at alkaline pH, and enables hydrophobic interaction C18 stationary phase through pairing with the acid functionality of the lipoic acid.


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[29,12]2--

[29,11]2-

Figure 1. Ag29(LA)12 mass spectrum (m/z 500 – 6000) acquired via direct infusion. Doubly charged anions with some fragmentation and moderate signal-to-noise is observed.


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Figure 2. LC-MS analysis of Ag29(LA)12 raw mixture sample. Black trace corresponds to the base peak chromatogram (m/z 100 – 6000). Orange and red traces correspond to extracted ion chromatograms (y-axis expanded) for components tentatively identified as dimer- and trimer-like species. LC conditions were 25 – 50% MP B over 20 minutes, followed by 5 min hold at 100% MP B. ESI time-of-flight mass spectra matching to the numbered Ag29(LA)12 isomers are shown inset in the Figure. LC peaks track with species as follows: (1, 2) Ag29(LA)12, 5.6 kDa; (3) Ag61(LA)24, 11.5 kDa (dimer); and (4) Ag93(LA)32, 17.5 kDa (trimer).


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Figure 3. LC method comparison showing the effect of gradient on retention and separation of Ag29(LA)12 isomers. (a) 35 – 50% MP B, (b) 30 – 50% MP B, (c) 25 – 50% MP B, (d) 20 – 50% MP B, (e) 10 – 50% MP B, and (f) 5 – 50% MP B.


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Figure 4. LC-UV trace (333 nm) for analysis of Ag29(LA)12 raw mixture sample. LC method conditions were 30 – 50% MP B over 20 minutes, followed by 5 min hold at 100% MP B. Overlaid UV-Vis spectra (300 – 500 nm) for each of the two Ag29(LA)12 isomers shown inset. Blue Trace: Peak 1; Red Trace: Peak 2.


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Figure 5. Representative in-source fragmentation spectrum of Ag29(LA)12. (Each of the two isobaric components fragments in virtually identical fashion.) (a) [Ag29(S2R)12H]2-, (b) [Ag29(S2R)11H]2- (c) [Ag29(S2R)10H]2-, (d) [Ag29(S2R)9SH]2-, (e) [Ag29(S2R)8S2H]2-, (f) [Ag29(S2R)7S3H]2-, (g) [Ag29(S2R)6S4H]2-, (h) [Ag29(S2R)5S5H]2-, (i) [Ag29(S2R)4S4H]2-, and (j) [Ag29(S2R)3S7H]2-.


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Label a b c d e f g h i j

Product ion [Ag29(S2R)12H]2[Ag29(S2R)11H]2[Ag29(S2R)10H]2[Ag29(S2R)9SH]2[Ag29(S2R)8S2H]2[Ag29(S2R)7S3H]2[Ag29(S2R)6S4H]2[Ag29(S2R)5S5H]2[Ag29(S2R)4S4H]2[Ag29(S2R)3S7H]2-

Valence 6 8 10 10 10 10 10 10 10 10

Table 1. Proposed product ion assignments and electron shell count for selected peaks observed in Figure 5. Electron count is determined according to the formula, ne = NAg - 2NS2R - 2NS - NH + z where ne is the number of valence shell electrons, NAu is the number of gold atoms in the cluster, NS2R is the number of thiolate ligands, NS is the number of free sulfur atoms, NH is the number of proton adduct ions, and z is the charge state of the ion.


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Scheme 2. Proposed in-source fragmentation processes: (a) neutral loss of lipoic acid moeity, (b) neutral loss of mercaptooctanoic acid moiety, where q = 1−7.


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ASSOCIATED CONTENT AUTHOR INFORMATION Supporting Information Supporting Information Available: LC-MS chromatograms acquired under non-ion-pairing conditions; LC-MS chromatograms acquired with acetonitrile in place of methanol; In-source fragmentation spectra for each of the two Ag29(LA)12 LC peaks; Positive mode LC-MS chromatogram, corresponding positive mode ESI mass spectra and related peak assignments.

Corresponding Authors *Robert L Whetten ([email protected]) *David M Black ([email protected]) Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding Sources The authors would like to acknowledge support from the National Institute on Minority Health and Health Disparities (G12MD007591), PREM: NSF PREM Grant #DMR 0934218; ‘‘Oxide and Metal Nanoparticles – The Interface Between Life Sciences and Physical Sciences”, WelchFoundation Grant AX-1857; “Fundamental Chemical Research on Larger Molecular NobleMetal Clusters” and the Texas Stars Fund. Notes The authors declare no competing financial interest. ABBREVIATIONS


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LA, lipoic acid; MPC, monolayer-protected cluster; TEA, triethylamine; HFIP, hexafluoroisopropanol; LC-MS, liquid chromatography mass spectrometry; ESI, electrospray ionization; ToF, time-of-flight.


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REFERENCES (1) Jin, R. Nanoscale 2015, 7, 1549-1565. (2) Schaaff, T. G.; Whetten, R. L. The Journal of Physical Chemistry B 2000, 104, 2630-2641. (3) Templeton, A. C.; Wuelfing, W. P.; Murray, R. W. Accounts of Chemical Research 2000, 33, 27-36. (4) Díez, I.; Pusa, M.; Kulmala, S.; Jiang, H.; Walther, A.; Goldmann, A. S.; Müller, A. H.; Ikkala, O.; Ras, R. H. Angewandte Chemie International Edition 2009, 48, 2122-2125. (5) Carducci, T. M.; Murray, R. W. Nanoelectrochemistry 2015, 73-124. (6) Li, G.; Jin, R. Accounts of chemical research 2013, 46, 1749-1758. (7) Yamazoe, S.; Koyasu, K.; Tsukuda, T. Accounts of chemical research 2013, 47, 816-824. (8) Agrachev, M.; Antonello, S.; Dainese, T.; Ruzzi, M.; Zoleo, A.; Aprà, E.; Govind, N.; Fortunelli, A.; Sementa, L.; Maran, F. ACS Omega 2017, 2, 2607-2617. (9) Zhu, M.; Aikens, C. M.; Hendrich, M. P.; Gupta, R.; Qian, H.; Schatz, G. C.; Jin, R. J. Am. Chem. Soc. 2009, 131, 2490-2492. (10) Kurashige, W.; Niihori, Y.; Sharma, S.; Negishi, Y. Coordination Chemistry Reviews 2016, 320-321, 238-250. (11) Ishida, Y.; Narita, K.; Yonezawa, T.; Whetten, R. L. The journal of physical chemistry letters 2016, 7, 3718-3722. (12) Brust, M.; Walker, M.; Bethell, D.; Schiffrin, D. J.; Whyman, R. J. Chem. Soc., Chem. Commun. 1994, 801-802. (13) Alvarez, M. M.; Khoury, J. T.; Schaaff, T. G.; Shafigullin, M. N.; Vezmar, I.; Whetten, R. L. The Journal of Physical Chemistry B 1997, 101, 3706-3712. (14) Morones, J. R.; Elechiguerra, J. L.; Camacho, A.; Holt, K.; Kouri, J. B.; Ramírez, J. T.; Yacaman, M. J. Nanotechnology 2005, 16, 2346. (15) Morones-Ramirez, J. R.; Winkler, J. A.; Spina, C. S.; Collins, J. J. Science translational medicine 2013, 5, 190ra181-190ra181. (16) Durán, N.; Durán, M.; de Jesus, M. B.; Seabra, A. B.; Fávaro, W. J.; Nakazato, G. Nanomedicine: Nanotechnology, Biology and Medicine 2016, 12, 789-799. (17) Franci, G.; Falanga, A.; Galdiero, S.; Palomba, L.; Rai, M.; Morelli, G.; Galdiero, M. Molecules 2015, 20, 8856-8874. (18) Natan, M.; Banin, E. FEMS Microbiol. Rev. 2017, 41, 302-322. (19) Niska, K.; Knap, N.; Kędzia, A.; Jaskiewicz, M.; Kamysz, W.; Inkielewicz-Stepniak, I. International journal of medical sciences 2016, 13, 772. (20) Azubel, M.; Koivisto, J.; Malola, S.; Bushnell, D.; Hura, G. L.; Koh, A. L.; Tsunoyama, H.; Tsukuda, T.; Pettersson, M.; Häkkinen, H. Science 2014, 345, 909-912. (21) Plascencia-Villa, G.; Demeler, B.; Whetten, R. L.; Griffith, W. P.; Alvarez, M.; Black, D. M.; José-Yacamán, M. The Journal of Physical Chemistry C 2016, 120, 8950-8958. (22) Black, D. M.; Bhattarai, N.; Whetten, R. L.; Bach, S. B. The Journal of Physical Chemistry A 2014, 118, 10679-10687. (23) Bhattarai, N.; Black, D. M.; Boppidi, S.; Khanal, S.; Bahena, D.; Tlahuice-Flores, A.; Bach, S.; Whetten, R. L.; Jose-Yacaman, M. The Journal of Physical Chemistry C 2015, 119, 1093510942. (24) Negishi, Y.; Nobusada, K.; Tsukuda, T. J. Am. Chem. Soc. 2005, 127, 5261-5270. (25) Knoppe, S.; Boudon, J.; Dolamic, I.; Dass, A.; Bürgi, T. Anal. Chem. 2011, 83, 5056-5061. (26) Qian, H.; Zhu, Y.; Jin, R. J. Am. Chem. Soc. 2010, 132, 4583-4585.


25


Page 26 of 28

(27) Ghosh, A.; Hassinen, J.; Pulkkinen, P.; Tenhu, H.; Ras, R. H.; Pradeep, T. Anal. Chem. 2014, 86, 12185-12190. (28) Jimenez, V. L.; Leopold, M. C.; Mazzitelli, C.; Jorgenson, J. W.; Murray, R. W. Anal. Chem. 2003, 75, 199-206. (29) Wolfe, R. L.; Murray, R. W. Anal. Chem. 2006, 78, 1167-1173. (30) Song, Y.; Jimenez, V.; McKinney, C.; Donkers, R.; Murray, R. W. Anal. Chem. 2003, 75, 5088-5096. (31) Choi, M. M.; Douglas, A. D.; Murray, R. W. Anal. Chem. 2006, 78, 2779-2785. (32) Negishi, Y.; Nakazaki, T.; Malola, S.; Takano, S.; Niihori, Y.; Kurashige, W.; Yamazoe, S.; Tsukuda, T.; Häkkinen, H. J. Am. Chem. Soc. 2015, 137, 1206-1212. (33) Negishi, Y.; Iwai, T.; Ide, M. Chemical Communications 2010, 46, 4713-4715. (34) Niihori, Y.; Matsuzaki, M.; Pradeep, T.; Negishi, Y. J. Am. Chem. Soc. 2013, 135, 49464949. (35) Niihori, Y.; Uchida, C.; Kurashige, W.; Negishi, Y. Physical Chemistry Chemical Physics 2016, 18, 4251-4265. (36) Knoppe, S.; Azoulay, R.; Dass, A.; Bürgi, T. J. Am. Chem. Soc. 2012, 134, 20302-20305. (37) Black, D. M.; Alvarez, M. M.; Yan, F.; Griffith, W. P.; Plascencia-Villa, G.; Bach, S. B.; Whetten, R. L. The Journal of Physical Chemistry C 2016, 121, 10851-10857. (38) Black, D. M.; Bhattarai, N.; Bach, S. B.; Whetten, R. L. J. Phys. Chem. Lett 2016, 7, 31993205. (39) Black, D. M.; Bach, S. B.; Whetten, R. L. Anal. Chem. 2016, 88, 5631-5636. (40) AbdulHalim, L. G.; Bootharaju, M. S.; Tang, Q.; Del Gobbo, S.; AbdulHalim, R. G.; Eddaoudi, M.; Jiang, D.-e.; Bakr, O. M. J. Am. Chem. Soc 2015, 137, 11970-11975. (41) Baksi, A.; Ghosh, A.; Mudedla, S. K.; Chakraborty, P.; Bhat, S.; Mondal, B.; Krishnadas, K.; Subramanian, V.; Pradeep, T. The Journal of Physical Chemistry C 2017, 121, 13421-13427. (42) Chakraborty, P.; Baksi, A.; Khatun, E.; Nag, A.; Ghosh, A.; Pradeep, T. The Journal of Physical Chemistry C 2017, 121, 10971-10981. (43) Kumar, J.; Kawai, T.; Nakashima, T. Chemical Communications 2017, 53, 1269-1272. (44) van der Linden, M.; Barendregt, A.; van Bunningen, A. J.; Chin, P. T.; Thies-Weesie, D.; de Groot, F. M.; Meijerink, A. Nanoscale 2016, 8, 19901-19909. (45) Russier-Antoine, I.; Bertorelle, F.; Hamouda, R.; Rayane, D.; Dugourd, P.; Sanader, Ž.; Bonačić-Koutecký, V.; Brevet, P.-F.; Antoine, R. Nanoscale 2016, 8, 2892-2898. (46) Lopez, P.; Lara, H. H.; Black, D. M.; Ramsower, H.; Alvarez, M. M.; Williams, T.; Demeler, B.; Lopez-Ribot, J. L.; Yacaman, M. J.; Whetten, R. L. 2017. (47) Black, D. M.; Crittenden, C. M.; Brodbelt, J. S.; Whetten, R. L. The Journal of Physical Chemistry Letters 2017, 8, 1283-1289. (48) Fountain, K. J.; Gilar, M.; Gebler, J. C. Rapid communications in mass spectrometry 2003, 17, 646-653. (49) Apffel, A.; Chakel, J. A.; Fischer, S.; Lichtenwalter, K.; Hancock, W. S. Anal. Chem. 1997, 69, 1320-1325. (50) Apffel, A.; Chakel, J. A.; Fischer, S.; Lichtenwalter, K.; Hancock, W. S. Journal of chromatography A 1997, 777, 3-21. (51) Niihori, Y.; Kikuchi, Y.; Shima, D.; Uchida, C.; Sharma, S.; Hossain, S.; Kurashige, W.; Negishi, Y. Ind. Eng. Chem. Res. 2017, 56, 1029-1035. (52) Baksi, A.; Harvey, S. R.; Natarajan, G.; Wysocki, V. H.; Pradeep, T. Chemical Communications 2016, 52, 3805-3808.


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TOC Graphic


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Liquid Chromatography Separation and Mass Spectrometry Detection

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