Anal. Chem. 2007, 79, 6215-6221
Sample-First Preparation: A Method for
Surface-Assisted Laser Desorption/Ionization
Time-of-Flight Mass Spectrometry Analysis of
Cyclic Oligosaccharides
Hsin-Pin Wu,† Chih-Lin Su,† Hui-Chiu Chang,‡, § and Wei-Lung Tseng*,†,§
Department of Chemistry, National Sun Yat-sen University, Kaohsiung, Taiwan, Graduate Institute of Medicine, College of
Medicine, Kaohsiung Medical University, Kaohsiung, Taiwan, and National Sun Yat-sen UniversitysKaohsiung Medical
University Joint Research Center, Kaohsiung, Taiwan
A new sample preparation method for the analysis of cyclic
oligosaccharides in surface-assisted laser desorption/
ionization mass spectrometry (SALDI-MS) is presented.
We call this new technique “sample first method”, in
which a sample is deposited first and then bare gold
nanoparticles (AuNPs), which serve as the SALDI matrixes, are added to the top of the sample layer. The use
of the sample first method offers significant advantages
for improving shot-to-shot reproducibility, enhancing the
ionization efficiency of the analyte, and reducing sample
preparation time as compared to the dried-droplet method,
wherein samples and bare AuNPs are mixed and dried
together. The relative standard deviation (RSD) values of
the signal intensity as calculated from 65 sample spots
was 25% when the sample first methods were applied to
the analysis of β-cyclodextrin. The results were more
homogeneous as compared to the outcome using drieddroplet preparation of AuNPs (RSD ) 66%) and 2,5dihydroxybenzoic acid (RSD ) 209%). We also found out
that the optimal concentration of AuNP for ionization
efficiency is 7.4 nM (4.52 × 1012 particles/mL) while the
lowest detectable concentration of cyclic oligosaccharides
through this approach is 0.25 µM. Except for the cyclic
oligosaccharide, the proposed method was also applied
to the analyses of other biological samples, including
neutral carbohydrate and steroid, aminothiols, and peptides as well as proteins.
An effective matrix-assisted laser desorption/ionization mass
spectrometry (MALDI-MS) analysis is strongly dependent on the
sample preparation.1-6 It is therefore not surprising that a wide
variety of methods and protocols for sample preparation have been
developed, such as the dried-droplet,1 fast evaporation,2 vacuum
* To whom correspondence should be addressed. E-mail: tsengwl@
mail.nsysu.edu.tw. Fax: 011-886-7-3684046.
†
National Sun Yat-sen University.
‡
Kaohsiung Medical University.
§
National Sun Yat-sen UniversitysKaohsiung Medical University Joint
Research Center.
(1) Karas, M.; Hillenkamp, F. Anal. Chem. 1988, 60, 2299-2301.
(2) Vorm, O.; Roepstorff, P.; Mann, M. Anal. Chem. 1994, 66, 3281-3287.
(3) Papac, D. I.; Wong, A.; Jones, A. J. S. Anal. Chem. 1996, 68, 3215-3223.
10.1021/ac070847e CCC: $37.00
Published on Web 07/14/2007
© 2007 American Chemical Society
drying,3 sandwich,4 two-layer,5 and even a three-layer method.6
So far, one of the easiest and most frequently used methods for
detection of peptides and proteins by MALDI-MS is the drieddroplet method. In this method, the solid matrix, such as 2,5dihydroxybenzoic acid (DHB) and sinapinic acid, is mixed with a
sample at an optimal ratio and then dried on the MALDI target
plate. Unfortunately, the inhomogeneous cocrystallization of the
analytes with the matrix requires sweet-spot searching and results
in poor shot-to-shot and sample-to-sample reproducibility.7 This
is the main factor that hampers the application of MALDI-MS for
quantitative measurements. Other factors, such as solvent composition, pH, and the presence of matrix additives can alter the
crystallization behavior and thus improve the quality of the
preparations.8,9 The method applied for sample preparation still
observes the sweet-spot effect. To overcome this problem, liquid
matrixes such as glycerol10 and ionic liquid11 have been developed
and successfully applied in the MALDI-MS analysis of highly
sulfated oligosaccharides, peptides, proteins, and poly(ethylene
glycol).12-14 Although the use of liquid matrixes exhibits higher
signal reproducibility, it is still limited to low mass resolution and
poor ionization efficiency.
Another possibility for improving the spot homogeneity of
analyte/matrix preparation is the use of nanomaterials such as
(4) (a) Whiteaker, J. R.; Fenselau, C. C.; Fetterolf, D.; Steele, D.; Wilson, D.
Anal. Chem. 2004, 76, 2836-2841. (b) Canelle, L.; Pionneau, C.; Marie,
A.; Bousquet, J.; Bigeard, J.; Lutomski, D.; Kadri, T.; Caron, M.; JoubertCaron, R. Rapid Commun. Mass Spectrom. 2004, 18, 2785-2794.
(5) (a) Zheng, J.; Li, N.; Ridyard, M.; Dai, H.; Robbins, S. M.; Li, L. J. Proteome
Res. 2005, 4, 1709-1716. (b) Dai, Y.; Whittal, R. M.; Li, L. Anal. Chem.
1999, 71, 1087-1091.
(6) Keller, B. O.; Li, L. J. Am. Soc. Mass Spectrom. 2006, 17, 780-785.
(7) Tholey, A.; Heinzle, E. Anal. Bioanal. Chem. 2006, 386, 24-37.
(8) Cohen, S. L.; Chait, B. T. Anal. Chem. 1996, 68, 31-37.
(9) (a) Distler, A. M.; Allison, J. Anal. Chem. 2001, 73, 5000-5003. (b) Kim,
Y.; Hurst, G. B.; Doktycz, M. J.; Buchanan, M. V. Anal. Chem. 2001, 73,
2617-2624.
(10) (a) Overberg, A.; Karas, M.; Bahr, U.; Kaufmann, R.; Hillenkamp, F. Rapid
Commun. Mass Spectrom. 1990, 4, 293-296. (b) Strupat, K.; Karas, M.;
Hillenkamp, F.; Eckerson, C.; Lottspeich, F. Anal. Chem. 1994, 66, 464470.
(11) Armstrong, D. W.; He, L.; Liu, Y.-S. Anal. Chem. 1999, 71, 3873-3876.
(12) Laremore, T. N.; Murugesan, S.; Park, T.-J.; Avci, F. Y.; Zagorevski, D. V.;
Linhardt, R. J. Anal. Chem. 2006, 78, 1774-1779.
(13) Li, Y. L.; Gross, M. L. J. Am. Soc. Mass Spectrom. 2004, 15, 1833-1837.
(14) Mank, M.; Stahl, B.; Boehm, G. Anal. Chem. 2004, 76, 2938-2950.
Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 6215
gold nanoparticles (AuNPs),15 silicon nanoparticles,16 and carbon
nanotubes.17 As compared with the conventional organic matrix,
the use of nanomaterials for the analysis of small molecules can
reduce matrix ion interference. This approach, which is known
as the surface-assisted laser desorption/ionization (SALDI)-MS,
offers the advantages of simple sample preparation, high surface
areas, flexibility in the sample deposition conditions. and independence from irradiation wavelength.18 Furthermore, the high
molar matrix-to-analyte ratio (∼107-109 analytes/particle) is
obtained using nanomaterials as the SALDI matrixes and thus
contributing to high efficient ionization. Small organic compounds,
peptides, and even proteins have all been detected with remarkable sensitivities using nanomaterials.15-19 For example, the
applicability of Nile red-adsorbed AuNPs as the SALDI matrixes
was successfully demonstrated in the analysis of aminothiols.20
Without enrichment, the limit of detection of glutathione obtained
by SALDI-MS is down to 1.0 µM. However, the amount of analytes
adsorbed on the particle surface is sensitive to the adsorption time.
We found that detection sensitivity dramatically decreases with
the decrease in contact time between analytes and particles. To
achieve a faster analysis, a new sample preparation method should
be developed for SALDI-MS.
In this work, we report the application of a sample first
preparation method for the analysis of cyclic oligosaccharides
when using bare AuNPs as the assisted matrix. In the sample
first method, the analyte is deposited first and then followed by
the bare AuNPs. The cyclic oligosaccharides, including R-, β-, and
γ-cycodextrin (CD), are difficult to ionize by MALDI-MS, but they
could be cationized very efficiently using bare AuNPs as matrixes
in combination with a sample first preparation method. As
compared with the dried-droplet and matrix first methods, a
sample first method not only provides high sensitivity for the
measurement of neutral carbohydrate but also improves the spot
homogeneity. This practical method was further validated by the
analyses of biological samples, including neutral carbohydrate,
neutral steroid, aminothiols, and peptides.
EXPERIMENTAL SECTION
Chemicals. R-CD, β-CD, γ-CD, ribose, glucose, raffinose,
maltose, testosterone, progesterone, corticosterone, cysteine,
homocysteine, glutathione, angiotensin I, substance P, and insulin
were all obtained from Sigma (St. Louis, MO). Citric acid,
hydrogen tetrachloroaurate(III) dehydrate, sodium borohydride,
and DHB were purchased from Aldrich (Milwaukee, WI). Ethanol
and ammonium hydroxide were obtained form Acros (Geel,
Belgium). When 150 mM citric acid was used to prepare citrate
buffer, NH4OH (25-30%) was used to adjust the pH to 4.0. Milli-Q
ultrapure water was used in all of experiments.
Apparatus. A double-beam UV-visible spectrophotometer
(Cintra 10e, GBC Scientific Equipment Pty Ltd., Dandenong,
(15) McLean, J. A.; Stumpo, K. A.; Russell, D. H. J. Am. Chem. Soc. 2005, 127,
5304-5305.
(16) (a) Wen, X.; Dagan, S.; Wysocki, V. H. Anal. Chem. 2007, 79, 434-444.
(b) Finkel, N. H.; Prevo, B. G.; Velev, O. D.; He, L. Anal. Chem. 2005, 77,
1088-1095.
(17) (a) Xu, S.; Li, Y.; Zou, H.; Qiu, J.; Guo, Z.; Guo, B. Anal. Chem. 2003, 75,
6191-6195. (b) Ren, S.-F.; Zhang, L.; Cheng, Z.-H.; Guo, Y.-L. J. Am. Soc.
Mass Spectrom. 2005, 16, 333-339.
(18) Sunner, J.; Dratz, E.; Chen, Y.-C. Anal. Chem. 1995, 67, 4335-4342.
(19) Chen, C.-T.; Chen, Y.-C. Anal. Chem. 2005, 77, 5912-5919.
(20) Huang, Y.-F.; Chang, H.-T. Anal. Chem. 2006, 78, 1485-1493.
6216
Analytical Chemistry, Vol. 79, No. 16, August 15, 2007
Victoria, Australia) was used to measure the absorbance of the
AuNPs. The homemade dark-field scattering system is made of
an Olympus IX70 inverted microscope (Tokyo, Japan), a DP70
digital camera (Olympus, Tokyo, Japan), and high-numericalaperture dark-field condenser (NA ) 1.2-1.4; U-DCW, Olympus).21 White light from a 100-W halogen lamp and a focusing
lens within a condenser are angled with respect to the objective
(40×; NAs ) 0.55) so that the illuminating light does not directly
enter the objective; thus, this arrangement results in a low
background. The scattering images for a AuNP-deposited glass
slide were observed using dark-field scattering microscopy. We
note that glass slide was used to replace a standard MALDI plate
because the inverted microscope required a transparent substrate.
The ImageJ program (http://rsb.info.nih.gov/ij/) was used to
analyze the images.
MS experiments were performed in the positive-ion mode on
a reflectron-type time-of-flight (TOF) mass spectrometer (Autoflex,
Bruker) equipped with a 3-m flight tube. Desorption/ionization
was obtained by using a 337-nm-diameter nitrogen laser with a
3-ns pulse width. The available accelerating voltages existed in
the range from +20 to -20 kV. To obtain good resolution and
signal-to-noise (S/N) ratios, the laser power was adjusted to
slightly above the threshold and each mass spectrum was
generated by averaging 500 laser pulses.
Synthesis of AuNPs with NaBH4. The bare (uncapped)
AuNPs were prepared by the chemical reduction of metal salt
precursor (hydrogen tetrachloroaurate) in a liquid phase.22 Typically, a 200-µL aliquot of 0.05 M aqueous HAuCl4 solution was
added to 50 mL of deionized water. Subsequently, 600 µL of freshly
prepared 0.05 M NaBH4 solution was added to the system,
whereby the solution gradually changed color to ruby red. We
note that these bare AuNPs are stable for a couple weeks because
small ions such as sodium, potassium, and chloride are absorbed
on the surface of bare AuNPs. The average size distribution of
the AuNPs is 12 ( 1.9 nm, which is confirmed by TEM
measurements (data not shown).23 To further determine the
concentration of the nanoparticles, we assumed that the reduction
from gold(III) to gold atoms was 100% complete.24 The concentration of the original bare AuNPs is ∼3.7 nM (2.26 × 1012 particles/
mL), which we denote in this study as 1×.
Preparation of Samples. Three sample preparation methods
for analysis of cyclic oligosaccharides were tested before SALDIMS measurements. Although the drying speed can be increased
by rising the temperature, fast evaporation of sample solution will
result in an increase in aggregation degree of AuNPs.25 Thus,
three sample preparation methods were all performed at room
temperature.
(a) Dried-Droplet cmethod. The cyclic oligosaccharides (500
µL) were added separately to 1× AuNP solutions (500 µL).
Immediately, the resulting mixtures (1 µL) were pipetted into a
stainless steel 384-well target (Bruker Daltonics) and dried in air.
We note that the drying time for the resulting mixtures is
∼30 min.
(21) Tseng, W.-L.; Lee, K.-H.; Chang, H.-T. Langmuir 2005, 21, 10676-10683.
(22) Gole, A.; Murphy, C. J. Chem. Mater. 2004, 16, 3633-3640.
(23) Su, C.-L.; Tseng, W.-L. Anal. Chem. 2007, 79, 1626-1633.
(24) Neiman, B.; Grushka, E.; Lev, O. Anal. Chem. 2001, 73, 5220-5227.
(25) (a) Sen, D.; Spalla, O.; Tache´, O.; Haltebourg, P.; Thill, A. Langmuir 2007,
23, 4296-4302. (b) Wang, W.-N.; Lenggoro, I. W.; Okuyama, K. J. Colloid
Interface Sci. 2005, 288, 423-431.
Figure 1. Mass spectra of β-CD obtained by using three sample preparation methods: (A) dried-droplet, (B) sample first, and (C) matrix first
method. The full scan mass spectrum was acquired from m/z 200 to 2000 (inset). The 1× bare AuNPs were used as the matrixes while the
sample concentration was 10 µM. The peaks at m/z 1157.95 and 1173.95 are assigned to the [β-CD + Na]+ and [β-CD + K]+, respectively. A
total of 500 pulsed laser shots were applied under a laser fluence of 60 µJ.
(b) Matrix First Method. The 1× AuNP solutions (1 µL) were
deposited first onto the sample plate and let dry. Then, l µL of
the cyclic oligosaccharides was deposited onto the first layer and
allowed to dry in air.
(c) Sample First Method. The cyclic oligosaccharides (1 µL)
were deposited first onto the sample plate and let dry in air. To
search for an optimal sample first method for SALDI-MS measurements, different concentrations (0.5-3.0×; 1 µL each) of AuNPs
solutions were deposited onto the first layer and allowed to dry
in air. We note that the analysis of carbohydrates, steroids,
aminothiols, peptide, and proteins was conducted by use of the
sample first method.
In the cases where DHB was used as a comatrix for MALDIMS analysis, DHB (1.0 µL) was mixed with the cyclic oligosaccharides (1.0 µL). Saturated matrix solution was prepared by
dissolving DHB in 50% ethanol. The resulting mixtures were then
pipetted into the wells of the steel plate and dried in air at room
temperature prior to MALDI-MS measurements.
RESULTS AND DISCUSSION
Comparison of Sample Preparation Methods for SALDIMS Analysis. Although a variety of sample preparation methods
have been proposed for MALDI-MS analysis,1-6 their impact on
SALDI-MS has not been carefully studied. Our initial investigations
using the dried-droplet methods revealed some inherent problems.21 In particular, the ionization efficiency is determined by
the amounts of analyte adsorbed on a particle. Thus, high
ionization efficiency was obtained when the contact time between
the analyte of interest and particles reached the adsorption
equilibrium time. For example, the adsorption equilibrium time
of neutral carbohydrate on the bare AuNP (12 ( 1.9 nm) was ∼2
h. On the other hand, we showed in an earlier publication21 that
the use of AuNPs as the SALDI matrixes provided high detection
sensitivity for small neutral carbohydrates but not oligosaccharides. The lower ionization efficiency for oligosaccharides can be
attributed to larger steric hindrance, which led to less adsorption
between analyte and nanoparticles. To speed up the analysis time
and improve the detection sensitivity in SALDI-MS, we tested
different sample preparation methods that include dried-droplet,
sample first, and matrix first methods. Figure 1A displays the
SALDI mass spectrum of β-CD obtained from the dried-droplet
method whereby the sample and bare AuNP (1×) are mixed and
immediately deposited on the sample target. As expected, the lowintensity ions at m/z 1157.95 were observed because only small
amounts of β-CD were adsorbed on the surface of AuNPs within
a short contact time. The peaks at m/z 1157.95 and 1173.95
correspond to [β-CD + Na]+ and [β-CD + K]+, respectively. No
MH+ signal obtained by SALDI-MS was due to the low proton
affinity of saccharides. By using the sample first method, a similar
mass spectrum was found but with a high intensity as shown in
Figure 1B. When the AuNPs are deposited on top of the sample
to form the second layer, they would be distributed on the entire
sample surface. On the basis of this phenomenon, the matrix-toanalyte ratio was relatively increased as compared to the drieddroplet method. More importantly, the resultant sample spots were
smooth and caused a significant improvement in spot homogeneity. Interestingly, the ion intensity of β-CD obtained by the sample
first method was still relatively high as compared to our previous
results using dried-droplet methods wherein the analytes were
equilibrated with bare AuNPs for 2 h. Inthe sample first method,
the upper layer of the AuNPs was irradiated with the laser beam
and resulted in an increase in the local temperature. The heat
was then transferred to the bottom layer of the analyte molecules
Analytical Chemistry, Vol. 79, No. 16, August 15, 2007
6217
Figure 2. Normalized [β-CD + Na]+ ion intensities obtained from 65 different sample spots. The samples are prepared by the sample first
(black squares) and dried-droplet (gray squares) methods, respectively. The RSD of the data series are presented as bar graphs; i.e., the black
and gray bars indicate the RSD values obtained using the sample first and dried-droplet methods, respectively. A total of 50 pulsed laser shots
were applied under a laser fluence of 60 µJ. The other conditions were the same as those in Figure 1.
and became well desorbed/ionized. By contrast, the analytes in
the dried-droplet methods adsorbed on the surface of the particles
were irradiated with the laser beam before the AuNPs. Thus, the
local temperature of the AuNPs was relatively lower and resulted
in poor ionization efficiency for β-CD. To support our hypothesis,
we first deposited the AuNPs on the sample stage, followed by
the samples (Figure 1C). Apparently, this preparation method was
not effective for ionization of β-CD.
To investigate possible improvements in shot-to-shot reproducibility using the sample first method, we compared it with the
dried-droplet method in SALDI-MS. Figure 2 shows that the use
of the sample first method decreased the variability of the signal
intensities of β-CD that were collected from 65 different sample
spots. The relative standard deviation (RSD) values of the signal
intensity were 25% for the sample first method and 66% for the
dried-droplet method. By contrast, these variations were greater
than 209% when using the dried-droplet preparation of DHB with
β-CD (Supporting information, Figure S1). The homogeneity of
the AuNP distribution was further investigated using the UVvis absorption spectrometer and dark-field microscopy when the
dried-droplet and sample first methods were conducted in the
analysis of β-CD. After the AuNPs were mixed with the analyte
in solution, the plasmon absorption of AuNPs exhibited a slight
red shift from 526 to 529 nm (Supporting Information, Figure S2).
This is because the resonance wavelength of the surface plasmon
in metallic nanoparticles is highly dependent on the surrounding
medium.26 However, when the mixture of the AuNPs and β-CD
was dried on the target, the aggregation of AuNPs occurred and
thus led to an increase in absorbance at longer wavelengths.
Compared with the sample first method, the plasmon band for
the dried-droplet method was considerably red-shifted and broadened (Supporting Information, Figure S2). On the other hand,
dark-field microscopy is a useful imaging technique for distinguishing individual and aggregated nanoparticles.27 As soon as
(26) McFarland, A. D.; Van Duyne, R. P. Nano Lett. 2003, 3, 1057-1062.
6218 Analytical Chemistry, Vol. 79, No. 16, August 15, 2007
Figure 3. Dark-field images of AuNPs obtained from (A) dried-droplet and (B) sample first methods. The 1× bare AuNPs were used as
matrixes, and the concentration of β-CD was 1 mM. Exposure times,
50 ms; scattering detection area using a 40× objective, 860 µm ×
610 µm corresponding to 4080 (horizontal) × 3072 (vertical) pixels.
the AuNPs were aggregated, the red shift of scattering would
occur, and hence, the aggregation of the AuNP was easily
Figure 4. Effect of AuNP concentration on the ionization efficiency of cyclic oligosaccharides: (A) 1×, (B) 1.5×, (C) 2×, (D) 2.5×, and (E) 3×.
The samples were prepared by the sample first method. The sample concentration was 2 µM. Peak identities: m/z 995.55, [R-CD + Na]+; m/z
1157.95, [β-CD + Na]+; m/z 1320.45, [γ-CD + Na]+. The other conditions were the same as those in Figure 1.
Figure 5. SALDI mass spectra of R- (0.25 µM), β- (0.25 µM), and γ-CD (0.5 µM) obtained using 2× AuNPs as the matrix. The samples were
prepared by the sample first method. Peak identities: m/z 995.55, [R-CD + Na]+; m/z 1011.55, [R-CD + K]+; m/z 1157.95, [β-CD + Na]+; m/z
1173.95, [β-CD + K]+; m/z 1320.45, [γ-CD + Na]+. The other conditions were the same as those in Figure 1.
recognized by conducting scattering measurements.28 Figure 3A
reveals that the AuNPs significantly aggregated using the drieddroplet method. The orange and red spots corresponded to the
scattering images of the AuNPs aggregates. It should be noted
(27) Xu, X.-H. N.; Brownlow, W. J.; Kyriacou, S. V.; Wan, Q.; Viola, J. J.
Biochemistry 2004, 43, 10400-10413.
(28) So
¨nnichsen, C.; Reinhard, B. M.; Liphardt, J.; Alivisatos, A. P. Nat. Biotechnol.
2005, 23, 741-745.
that the orange spot came from an aggregate of more than two
particles. In comparison, Figure 3B shows that some spots
displayed a green color, which corresponded to the scattering
images of single AuNPs.29 It was suggested that an increase in
homogeneity of the analyte could be achieved by using the
sample first method (Figure 3B). These finding support the
(29) Orendorff, C. J.; Sau, T. K.; Murphy, C. J. Small 2006, 2, 636-639.
Analytical Chemistry, Vol. 79, No. 16, August 15, 2007
6219
notion that the use of the sample first method offers a better
reproducibility as a result of the improvement in the sample
homogeneity.
Effect of AuNP Concentration. When the AuNPs were used
as the SALDI matrixes in the dried-droplet method, the ionization
efficiency of the analyte was greatly affected by the particle
concentration. Thus, the effect of the AuNP concentrations on
the signal intensities of cyclic oligosaccharides was also studied
in the sample first method. Figure 4 shows that the signal
intensities of a mixture comprising equal concentrations (2 µM)
of the R-, β-, and γ-CD were maximized while the AuNP
concentration increased by up to 2×. The enhancement was likely
due to the increase in the number of the deposited AuNPs on top
of the sample. Under this condition, we ensured that the entire
sample surface was covered with AuNPs. However, a decrease in
the signal was observed above 2× the level of concentration. This
finding indicates that a thicker layer was formed when a higher
concentration of AuNPs was deposited. This phenomenon would
increase the isolated AuNPs and thereby result in the decrease
of the local temperature. In comparison with the use of 1× AuNPs,
the optimization of the concentration of AuNPs provided more
than a 3-fold enhancement in the S/N ratio for cyclic oligosaccharides. Figure 5 shows the SALDI mass spectrum of a
mixture containing R- (0.25 µM), β-(0.25 µM), and γ-CD (0.5 µM).
However, no signals were identifiable when their concentrations
were lowered to 0.1 µM, which indicated that the detection limits
were somewhere between 100 and 250 fmol. By contrast, this
method was 105 times more sensitive for the analysis of cyclic
oligosaccharides than with the use of DHB as MALDI matrixes.30
Although the ionization efficiency of MALDI-MS can be further
enhanced by reacting cyclic oligosaccharides with succinic
anhydride, a long reaction time (24 h) is unduly required.30
Analysis of Different Biomolecules. The successful analysis
of neutral, cyclic oligosaccharides, as described earlier, motivated
us to apply the sample first method to the analysis of other
biological compounds. The neutral carbohydrates and steroid,
aminothiols, and peptides were tested by SALDI-MS in combination with the sample first method. Figure 6A displays the SALDI
mass spectrum of the carbohydrate mixture (5 µM). The peaks
that appear at m/z 172.86, 202.91, 364.74, and 526.76 corresponded
to the sodiated molecular ions of ribose, glucose, maltose, and
raffinose, respectively. In contrast to the SALDI-MS spectra
obtained with the dried-droplet method, the use of the sample
first method provided better sensitivity (except for glucose) and
decreased the analysis time. To further explore the capability of
the sample first method to analyze more hydrophobic compounds,
a mixture of steroids (50 µM) was chosen in this study. It should
be noted that steroids are ionized poorly by both MALDI and
electrospray ionization MS.31 The successful ionization of neutral
steroids is shown in Figure 6B. The peaks appearing at m/z
311.13, 337.14, and 369.15 were the sodiated molecular ions of
testosterone, progesterone, and corticosterone, respectively. We
were also interested in determining the thiol-containing amino
acids, which were successfully ionized by using Nile red-absorbed
(30) Ahn, Y. H.; Yoo, J. S.; Kim, S. H. Anal. Sci. 1999, 15, 53-56.
(31) (a) Wang, Y.; Hornshaw, M.; Alvelius, G.; Bodin, K.; Liu, S.; Sjo¨vall, J.;
Griffiths, W. J. Anal. Chem. 2006, 78, 164-173. (b) Griffiths, W. J.; Liu, S.;
Alvelius, G.; Sjo
¨vall, J. Rapid Commun. Mass Spectrom. 2003, 17, 924935.
6220 Analytical Chemistry, Vol. 79, No. 16, August 15, 2007
Figure 6. Sample first method for SALDI-MS analysis of biomolecules: (A) neutral carbohydrates (5 µM), (B) neutral steroids (50
µM), and (C) aminothiols (200 µM). Peak identities: (A) m/z 172.86,
[ribose + Na]+; m/z 202.91, [glucose + Na]+; m/z 364.74, [maltose
+ Na]+; m/z 526.76, [raffinose + Na]+. (B) m/z 311.13, [testosterone
+ Na]+; m/z 337.14, [progesterone + Na]+; m/z 369.15, [corticosteron
+ Na]+. (C) m/z 144.00, [cysteine + Na]+; m/z 158.00, [homocysteine
+ Na]+; m/z 329.98, [glutathione + Na]+. The sample concentration
was 10 µM while the other conditions were the same as those in
Figure 5.
AuNPs as the SALDI matrixes.20 Figure 6C displays the SALDI
mass spectra of a mixture containing cysteine, homocysteine, and
glutathione. It was suggested that the sample first method could
be directly used for the detection of aminothiols without the need
for contact time between aminothiols and AuNPs. Since the
successful implementation of SALDI-MS for the analysis of small
biomolecules was well established, we also extend this method
for peptides and proteins. Panels A and B in Figure 7 show that
substance P and insulin were successfully ionized by using AuNPs
as the SALDI matrixes. It should be noted that our proposed
method cannot be applied to the analysis of angiotensin I, which
is the standard peptide for MALDI-MS. It is believed that the
ionization of angiotensin I can be achieved by searching the
appropriate sample solution. The peaks that appeared at m/z
1370.64 and 5779.94 corresponded to [substance P + Na]+ and
[insulin + 2Na]+, respectively. The peak shape and ionization
efficiency of insulin were further improved when the sample was
prepared in a citrate buffer at pH 4.0 (Supporting Information,
Figure S3).32 However, difficulties were encountered in the
(32) Chen, W.-Y.; Chen, Y.-C. Anal. Bioanal. Chem. 2006, 386, 699704.
Figure 7. Sample first method for SALDI-MS analysis of (A)
substance P (10 µM) and (B) insulin (100 µM). Peak identities: (A)
m/z 1370.64, [substance P + Na]+; (B) m/z 5779.94, [insulin + 2Na]+
The other conditions were the same as those in Figure 5.
analysis of cytochrome c (12 300 Da), which indicated that the
upper detectable mass limit was ∼6000 Da. Moreover, the
detection sensitivity decreased dramatically with increase in mass.
The same phenomena mentioned above were also reported by
Hillenkamp’s group.33
CONCLUSIONS
The sample first method for SALDI-MS analysis of neutral and
charged biomolecules using bare AuNPs provides a number of
(33) Schurenberg, M.; Dreisewerd, K.; Hillenkamp, F. Anal. Chem. 1999, 71,
221-229.
(34) Harvey, D. J. Mass Spectrom. Rev. 1999, 18, 349-351.
(35) Stubiger, G.; Belgacem, O. Anal. Chem. 2007, 79, 3206-3213.
(36) Yu, Y. Q.; Fournier, J.; Gilar, M.; Gebler, J. C. Anal. Chem. 2007, 79, 17311738.
(37) Zhao, J.; Qiu, W.; Simeone, D. M.; Lubman, D. M. J. Proteome Res. 2007,
6, 1126-1138.
distinctive advantages. First, it allows the sensitive detection of
neutral oligosaccharides and steroid in their underivatized form
wherein the detection limits for R-, β-, and γ-CD are in the highfemtomole levels. Second, this method is simple and time-efficient
when nanomaterials are used as the SALDI matrixes. The analytes
can be detected directly without any adsorption steps. Third,
reproducibility of signal intensities is appreciably improved by the
sample first method mainly because of the improvement in the
spot homogeneity. Last, it can be applied in the analysis of peptides
and proteins, whose the upper detectable mass limit was ∼ 6000
Da. On the basis of these advantages, we believe that the proposed
methods can be applied not only to the analysis of many types of
carbohydrate and carbohydrate-containing compounds34 but also
in the detection of other neutral biomolecules, such as steroids31
and lipids.35 For example, we should be able to use the proposed
method for identifying glycosylation sites of proteins after the
appropriate enzyme digestion.36 Also, the method can be helpful
in the comparative studies of the carbohydrate chains of glycoproteins produced by malignant cells and normal cells.37
ACKNOWLEDGMENT
We thank the National Science Council (NSC 95-2113-M-110020-) of Taiwan and Aim for the Top University Plan, Ministry of
Education, Taiwan for the financial support of this study. We thank
Professor J. Shiea for allowing us to use the MALDI-TOF
instrument.
SUPPORTING INFORMATION AVAILABLE
Additional information as noted in text. This material is
available free of charge via the Internet at http://pubs.acs.org.
Received for review April 26, 2007. Accepted June 13,
2007.
AC070847E
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