Anal. Chem. 2010, 82, 7096–7102
In Vivo Monitoring of Quantum Dots in the Extracellular Space Using Push-Pull Perfusion Sampling, Online In-Tube Solid Phase Extraction, and Inductively Coupled Plasma Mass Spectrometry C. K. Su,† C. W. Huang,† C. S. Yang,§ Y. J. Wang,§ and Y. C. Sun*,†,‡ Department of Biomedical Engineering and Environmental Sciences and Nuclear Science and Technology Development Center, National Tsing-Hua University, Hsinchu, Taiwan, and Center for Nanomedicine Research, National Health Research Institutes, Zhunan, Miaoli, Taiwan To monitor the dynamic changes of extracellular quantum dots (QDs) in vivo in the livers of anesthetized rats, we developed an automatic online analytical system comprising push-pull perfusion (PPP) sampling, the established in-tube solid phase extraction (SPE) procedure, and inductively coupled plasma mass spectrometry (ICPMS). The method takes advantage of the retention of QDs onto the interior surface of a polytetrafluoroethylene (PTFE) tube as a means of extracting the QDs from complicated push-pull perfusates. For the injected QDs present in the liver extracellular fluid (ECF) at low picomolar levels, a temporal resolution of 10 min was required to collect sufficient amounts of QDs to meet the sensitivity requirements of the ICPMS system. To the best of our knowledge, this study is the first to exploit the PPP technique for the collection of QDs from living animals and PTFE tubing as a SPE adsorbent for the online extraction of QDs and the removal of biological matrix prior to ICPMS analysis of cadmium-containing inorganic nanocrystal. We confirmed the analytical reliability of this method from measurements of the spike recoveries of saline samples; in addition, we demonstrated the systems’ applicability through in vivo monitoring of the time-dependent concentration profile of liver extracellular QDs in living rats after intravenous administration. The emergence and progression of nanomaterial-based agents or fluorophores for biomedical imaging applications have spurred investigations into the interactions between nanosized materials (NMs) and biological systems. Among these suitable candidate NMs, cadmium-containing quantum dots (QDs) are widely used nanocrystals for biomedical sensing and imaging applications,1-3 * To whom correspondence should be addressed. Fax: +886-3-5723883. Tel.: +886-3-5727309. E-mail:
[email protected]. † Department of Biomedical Engineering and Environmental Sciences, National Tsing-Hua University. ‡ Nuclear Science and Technology Development Center, National Tsing-Hua University. § National Health Research Institutes. (1) Bruchez, M., Jr.; Moronne, M.; Gin, P.; Weiss, S.; Alivisatos, A. P. Science 1998, 281, 2013–2016.
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due mainly to their unique and tunable optical properties.4 To enable more precise quantitative biology, biomedical research seeks to use bioconjugated QDs to target specific biological molecules and probe nanoenvironments in the “right place at the right time.” From pharmacological and toxicological standpoints, however, before QDs gain approval and enter practical use, we must have the thorough understanding of their pharmaco-/toxicokinetic characteristics and dynamic biodistribution.5-7 Biological studies of QDs have revealed that their cellular responses and biodistributions are dependent on their chemical compositions, sizes, shapes, and surface modifications.8,9 In addition, diffusion or active intracellular transport within the organ and/or cell is the typical path for the inter- and intracellular transport of QDs in living organisms.10-12 To understand the transfer kinetics of QDs, information is required on the responses of living systems toward the presence of extracellular QDs of various sizes, shapes, chemical compositions, and degrees of surface modification, as well as the temporal fate of QDs that are subject to translocation and degradation processes.11,13 Nevertheless, because of the lack of adequate methods for in vivo monitoring of the transfer kinetics of QDs in the extracellular (2) Chan, W. C. W.; Nie, S. Science 1998, 281, 2016–2018. (3) Gao, X.; Cui, Y.; Levenson, R. M.; Chung, L. W. K.; Nie, S. Nat. Biotechnol. 2004, 22, 969–976. (4) Resch-Genger, U.; Grabolle, M.; Cavaliere-Jaricot, S.; Nitschke, R.; Nann, T. Nat. Methods 2008, 5, 763–775. (5) Nel, A.; Xia, T.; Maodler, L.; Li, N. Science 2006, 311, 622–627. (6) Colvin, V. L. Nat. Biotechnol. 2003, 21, 1166–1170. (7) Fischer, H. C.; Chan, W. C. W. Curr. Opin. Biotechnol. 2007, 18, 565– 571. (8) Geys, J.; Nemmar, A.; Verbeken, E.; Smolders, E.; Ratoi, M.; Hoylaerts, M. F.; Nemery, B.; Hoet, P. H. M. Environ. Health Perspect. 2008, 116, 1607–1613. (9) Simon-Deckers, A.; Loo, S.; Mayne-L’hermite, M.; Herlin-Boime, N.; Menguy, N.; Reynaud, C.; Gouget, B.; Carrie´re, M. Environ. Sci. Technol. 2009, 43, 8423–8429. (10) Gao, X.; Wang, T.; Wu, B.; Chen, J.; Chen, J.; Yue, Y.; Dai, N.; Chen, H.; Jiang, X. Biochem. Biophys. Res. Commun. 2008, 377, 35–40. (11) Scientific Committee on Emerging and Newly Identified Health Risks (SCENIHR): Opinion on-The appropriateness of existing methodologies to asses the potential risks associated with engineered and adventitious products of nanotechnologies. 28-29th Sept. 2005, SCENIHR/002/05. (12) Hess, H.; Tseng, Y. ACS Nano 2007, 1, 390–392. (13) Hild, W. A.; Breunig, M.; Goepferich, A. Eur. J. Pharm. Biopharm. 2008, 68, 153–168. 10.1021/ac100167v 2010 American Chemical Society Published on Web 08/13/2010
space of target organs, the link between QD exposure and human risk remains unclear; exploring the physiopathological roles of various QDs/NMs remains a demanding challenge for the development of nanotechnology. To satisfy the increasing demand for information regarding the pharmaco-/toxico-kinetic characteristics of QDs in experimental animals, fluorescence measurements3,14-16 have become the preliminary strategy for experimentally determining their disposition kinetics. The major drawbacks of this technique for in vivo imaging are that it only provided the relative quantification of fluorescence intensity and was limited by the penetration depth of the emitted light itself. Moreover, photon attenuation is not linear as a function of depth and optical heterogeneity of tissue, which could obscure the results of quantification.17 To simplify the data acquisition process, radioactive tracer techniques18-20 and elemental analysis techniques8,15,16 have been applied as alternatives to optical imaging when acquiring quantitative biodistribution and monitoring the tissue kinetics of QDs after exposure. Labeling with radioactive 64Cu, for example, allows the ready monitoring of the in vivo distributions and time-course relationships of QDs.18,19 Although this technique is sensitive and can yield desirable biodistribution data, modification of the radionuclide on the QDs might change the surface properties (e.g., surface hydrophilicity or charge density), thereby, influencing the biodistribution. Furthermore, the conjugated radionuclide or crosslinker molecules might detach in complicated biological environments, thereby distorting the biodistribution pattern and limiting the diagnostic value of the radiolabeled QDs.21 At present, batchwise elemental analysis remains a userfriendly technique for generating quantitative static data.8,15,16 After sacrificing animals at period lengths of time after their exposure to QDs, the organs and tissues of interest are collected and the bulk concentration of the tested QDs are determined using appropriate sample preparation and instrumental measurement. However, because of the involvement of the autopsy sampling technique and vigorous sample digestion procedure, the main disadvantage of this method is that it has no capability of providing the details of kinetic distribution of QDs in the target organs and tissues. This paper describes a facile technique for characterizing the biokinetics of extracellular QDs in the livers of living rats after intravenous administration. To characterize and monitor transfer kinetics of extracellular QDs in vivo, we first used push-pull perfusion sampling to continuously collect QD-containing extra(14) Choi, H. S.; Liu, W.; Misra, P.; Tanaka, E.; Zimmer, J. P.; Ipe, B. I.; Bawendi, M. G.; Frangioni, J. V. Nat. Biotechnol. 2007, 25, 1165–1170. (15) Yang, R. S. H.; Chang, L. W.; Wu, J.; Tsai, M.; Wang, H.; Kuo, Y.; Yeh, T.; Yang, C. S.; Lin, P. Environ. Health Perspect. 2007, 115, 1339–1343. (16) Fischer, H. C.; Liu, L. C.; Pang, K. S.; Chan, W. C. W. Adv. Funct. Mater. 2006, 16, 1299–1305. (17) Ntziachristos, V.; Bremer, C.; Weissleder, R. Nat. Biotechnol. 2005, 23, 313–320. (18) Schipper, M. L.; Iyer, G.; Koh, A. L.; Cheng, Z.; Ebenstein, Y.; Aharoni, A.; Keren, S.; Bentolila, L. A.; Li, J.; Rao, J.; Chen, X.; Banin, U.; Wu, A. M.; Sinclair, R.; Weiss, S.; Gambhir, S. S. Small 2009, 5, 126–134. (19) Schipper, M. L.; Cheng, Z.; Lee, S.; Bentolila, L. A.; Iyer, G.; Rao, J.; Chen, X.; Wu, A. M.; Weiss, S.; Gambhir, S. S. J. Nucl. Med. 2007, 48, 1511– 1518. (20) Woodward, J. D.; Kennel, S. J.; Mirzadeh, S.; Dai, S.; Wall, J. S.; Richey, T.; Avenell, J.; Rondinone, A. J. Nanotechnology 2007, 18, 175103. (21) Marquis, B. J.; Love, S. A.; Braun, K. L.; Haynes, C. L. Analyst 2009, 134, 425–439.
cellular fluid (ECF) from a living animal. In view of the severe interference resulting from the presence of large numbers of blood cells, proteins, and dissolved salts in the push-pull perfusates, it was necessary for us to develop an effective pretreatment process to remove the complicated matrix and enrich the trace amounts of QDs to obtain reliable inductively coupled plasma mass spectrometry (ICPMS) data. The most frequently encountered separation procedures for NMs include diafiltration,22 capillary electrophoresis,23 size-exclusion chromatography,24,25 hydrodynamic chromatography,26 and field flow fractionation.27 Nevertheless, the strong interactions between NMs and the separation materials can complicate these elution processes and decrease their recoveries.25,28 In addition, the concomitants described above could also adsorb irreversibly onto the separation materials and deteriorate their long-term stability for the separation of NMs of interest.29 During the past decade, Fang et al.30 developed a unique online preconcentration system based on the retention of metal precipitates on the inner walls of a knotted reactor (KR) prepared using a polytetrafluoroethylene (PTFE) tube. Because of its simplicity and high cost-effectiveness, this novel procedure has been applied widely to the determination of trace elements in biological and environmental samples.31,32 Because the use of open tubular reactors, such as KRs, eliminates the drawbacks associated with flow resistance during packed-bed solid phase extraction (SPE) and the demand for high pressure pumps, we suspected that a PTFE tube would allow online preconcentration of QDs in samples featuring high salt and protein contents. To the best of our knowledge, PTFE tubing has never previously been incorporated into an online separation system for the SPE of QDs, even though it might lead to simpler and cleaner online separation systems exhibiting sample enrichment and separation capability. In this study, we developed a selective and sensitive hyphenated system for continuous determination of trace QDs in push-pull perfusates, employing a separation process that exploits the hydrophobic interactions between PTFE and QDs in conjunction with online ICPMS determination. This procedure does not require the use of any other reagents for the adsorption procedure nor are cleaning or postconditioning procedures necessary for the separation of QDs from the push-pull perfusates. After optimizing the operating conditions for the separation process, we arranged a Micromist nebulizer in series at the interface between the intube PTFE SPE device and the ICPMS system to establish a simple, rapid, and sensitive online push-pull perfusion (PPP) intube PTFE SPE-ICPMS hyphenated system to facilitate the in vivo (22) Sweeney, S. F.; Woehrle, G. H.; Hutchison, J. E. J. Am. Chem. Soc. 2006, 128, 3190–3197. (23) Surugau, N.; Urban, P. L. J. Sep. Sci. 2009, 32, 1889–1906. (24) Al-Somali, A. M.; Krueger, K. M.; Falkner, J. C.; Colvin, V. L. Anal. Chem. 2004, 76, 5903–5910. (25) Krueger, K. M.; Al-Somali, A. M.; Falkner, J. C.; Colvin, V. L. Anal. Chem. 2005, 77, 3511–3515. (26) Tiede, K.; Boxall, A. B. A.; Tiede, D.; Tear, S. P.; Davide, H.; Lewis, J. J. Anal. At. Spectrom. 2009, 24, 964–972. (27) Chun, J.; Fagan, J. A.; Hobbie, E. K.; Bauer, B. J. Anal. Chem. 2008, 80, 2514–2523. (28) Fischer, C.; Siebrands, T. J. Chromatogr., A 1995, 707, 189–197. (29) Misˇl’anova´, C.; Hutta, M. J. Chromatogr., B 2003, 797, 91–109. (30) Fang, Z.; Xu, S.; Dong, Z. L.; Li, W. Talanta 1994, 41, 2165–2172. (31) Yan, X. P.; Jiang, Y. TrACsTrends Anal. Chem. 2001, 20, 552–562. (32) Cerutti, S.; Martinez, L. D.; Wuilloud, R. G. Appl. Spectrosc. Rev. 2005, 40, 71–101.
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Figure 1. Schematic representation of the PPP in-tube PTFE SPE-ICPMS system for online extraction and monitoring of QDs in rat liver. VA, VC: Six-port rotary valves; VB: eight-port rotary valves; P1, P2, P3: peristaltic microdialysis pump; P4: conventional peristaltic pump.
determination of the biokinetics of QDs through the extracellular space in living rats. EXPERIMENTAL SECTION Chemicals and Reagents. The QDs used in these experiments were purchased from Invitrogen (Eugene, OR) as Qtracker 705 nontargeted QDs (QD705), which are cadmium/selenium/ tellurium (Cd/Se/Te)-based nanocrystals presenting an amphiphilic methoxy-PEG-5000 surface coating. According to the manufacturer’s instruction, its diameter and molecular weight are about 13 nm and 1.5 × 106 g mole-1, respectively. Phosphate-buffered saline (PBS; P3813, Sigma) solution containing heparin sodium (10 IU mL-1; B. Braun, Melsungen AG, Germany) was prepared from high-purity water. The eluent was prepared by dissolving nitric acid (20 mL, for trace metal analysis; J.T. Baker, NJ) in high-purity water (1 L). The infusion solution [0.9% NaCl (w/v) and anticoagulant (10 IU mL-1)] was prepared by dissolving sodium chloride (ultrapure grade, E. Merck, Darmstadt, Germany) and heparin sodium in highpurity water. Push-Pull Perfusion Sampling. Using a procedure for the construction of the modified probe similar to those reported previously,33 a concentric probe was developed to allow the direct infusion of saline solution to the desired sampling region via an outer infusion polyurethane tubing (o.d./i.d., 770/580 µm) and simultaneous sample collection via an inner withdraw polyimide tubing (o.d./i.d., 380/280 µm). The flow rate required to pull the sample from the sampling site was not set as an experimental variable but was adjusted daily to maintain a withdrawing flow rate equal to the infusion flow rate. Apparatus. The PPP in-tube PTFE SPE-ICPMS hyphenation system (Figure 1) consisted of three main parts: PPP sampling, (33) Kottegoda, S.; Shaik, I.; Shippy, S. A. J. Neurosci. Methods 2002, 121, 93– 101.
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an SPE device for QDs, and an ICPMS analysis system. The online PPP sampling system contained two peristaltic microdialysis pumps (MAB 20, Microbiotech/se AB, Stockholm, Sweden) and a homemade push-pull probe. The connections of the microdialysis pump to the inlet of the push-pull probe and the outlet to a mixing-tee (CM1XPK, Valco, Lucerne, Switzerland) were both formed using fluorinated ethylene polypropylene (FEP) tubing (internal volume: 1.2 µL per 100 mm length; length: 15 cm; i.d.: 0.12 mm; CMA, Stockholm, Sweden). The perfused sample was online-mixed with PBS solution, which was carried by another microdialysis pump; the mixture was delivered to a six-port valve (C2-2348D, Valco, Lucerne, Switzerland) through a 1.5 cm long, 0.007 in. i.d. piece of PTFE tubing. To reduce blood coagulation and the possibility of hydrodynamic impedance in the PTFE tube caused by clots, the PBS solution was heparinized in advance. The online flow injection pretreatment system for QDs comprised two six-port valves and an eight-port valve (C22Z-3186E, Valco, Lucerne, Switzerland). All the valves were programmed and controlled by a personal computer through a serial valve interface (SIV-110, Valco, Lucerne, Switzerland). The operational sequence of the online pretreatment system is given in Table 1. The connections and conduits were PTFE connecting tubes. The in-tube PTFE SPE device consisted of a tract of PTFE tubing (80 cm long × 0.007 in. i.d.; Alltech, Virginia). A conventional peristaltic pump was employed as a conduit for the samples and eluent (2% HNO3). The ICP mass spectrometer was an Agilent 7500a system (Agilent, CA). A Micromist nebulizer (AR35-1-EM04EX, Glass Expansion, Victoria, Australia) was fitted to a Scott-type quartz double-pass spray chamber. Because of possible memory effects and long washout times during the elution procedure, the QDs were quantified only in terms of the elution peak areas of cadmium ions. The instrumental operating conditions selected for optimal sensitivity and low background noise are presented in Table 2.
Table 1. Flow Injection Program Used for the Online In-Tube PTFE SPE-ICPMS Hyphenated System step 1
2 3 4
valve position
function
medium pumped
flow rate, µL min-1
push-pull perfusate
10
buffer
10
80
buffered sample
20
80
HNO3
400
20
H 2O
400
time, s
A: load fill heparinized PBS-buffered sample B: load into PTFE tubing C: load A: injection remove waste B: load C: load A: injection detach QDs and B: injection deliver to ICPMS C: load A: injection replace HNO3 with water carrier stream B: injection C: injection
420
Table 2. Operating Conditions for the Established PPP In-Tube PTFE SPE-ICPMS Hyphenated System ICPMS ICPMS spectrometer Ar gas flow rates outer auxiliary nebulizer make-up plasma forward power sampling cone skimmer cone
Agilent 7500a 15 L min-1 0.9 L min-1 1.03 L min-1 0.12 L min-1 1500 W Ni, 1 mm orifice Ni, 0.4 mm orifice
In-Tube Extraction tube material length of tube inner diameter of tube buffer solution for mixing sample loading eluent elution flow rate capacity
PTFE 80 cm 0.007 in. PBS solution (pH 7.4) containing heparin (10 IU mL-1) 20 µL min-1 2% HNO3 400 µL min-1 1.0 fmole QD705 cm-2
PPP sampling infusion solution infusion flow rate sampling time sampling frequency
0.9% NaCl (pH 7.4) 10 µL min-1 420 s 6 h-1
In Vivo Experiments. Adult male Sprague-Dawley rats (200-250 g) were obtained from the Laboratory Animal Center of the National Science Council of the Republic of China (Taipei, Taiwan). These animals, which were specifically pathogen-free, were acclimatized to their environmentally controlled quarters (25 °C; 12 h light/12 h dark cycle) for at least 3 days before experimentation and then fasted overnight prior to sacrifice. The rats were fed a standard diet and water and were treated under the regulations of the “Principles of Laboratory Animal Care” (NIH publication no. 86-23, revised, 1985). The study was approved by the committee of experimental animals of National Tsing-Hua University. To estimate the concentrations of QDs in vivo, it was necessary to determine the daily recovery of PPP sampling, which was accomplished by placing the probe in a Ringer’s solution of a
known concentration of QDs and perfusing at 10 µL min-1. The perfused sample was online mixed with a stream of PBS solution with the same flow rate and then introduced directly into an 80 cm long PTFE tube for the extraction of QDs and removal of the blood matrix. After removing the waste, the QDs adsorbed on the interior surface of the PTFE tube were dissolved using 2% (v/v) HNO3 and transported to the ICPMS system using another carrier stream (flow rate: 400 µL min-1) for final quantification. On the basis of the proportions of Cd, Se, and Te in QD705, the highest sensitivity was obtained when acquiring the ion intensities at m/z 112 (Cd) and 114 (Cd) to set up the calibration curves for quantification of the QDs. The rats were initially anesthetized with urethane (1600 mg kg-1 body weight, intraperitoneal injection); they remained anesthetized throughout the experimental period. A middle laparotomy was performed, and the liver hilum was exposed. The push-pull probe, which was perfused with saline solution (flow rate: 10 µL min-1), was implanted; after 1 h, a dosage of 200 pmole QD705 kg-1 body weight was intravenously injected into the rat and the level of the QDs was monitored continuously every 10 min. Histological Examination Following Sampling. At the conclusion of experiment, rats were euthanized. A liver tissue containing sampling site was rapidly harvested and placed in 10% formalin solution. After that, this tissue was embedded in paraffin. Sections cut on a microtome at 4 µm thickness were stained with hematoxylin and eosin (H and E) for further analysis by a pathologist. RESULTS AND DISCUSSION PTFE-QD705 Interaction. In this study, we adopted commercial PTFE tubes for initial online extraction of QDs from highsalt content push-pull perfusates collected from the extracellular space of rat livers. To investigate the retention behavior of QDs toward the interior wall of the PTFE tube, we mixed QD705, having surfaces covered with poly(ethylene glycol) (PEG) chains, individually with n-hexane, diethyl ether, ethanol, and water and then transported the mixtures into the PTFE tube to examine the effect of the polarity of the reaction environment on the extraction efficiency. The presence of nonpolar solvents (n-hexane, diethyl ether) led to negligible retention of the QDs on the interior wall of the PTFE tube. In contrast, nearly 100% of the QDs dispersed in the polar solvents (water, ethanol) were retained on the interior wall of the PTFE tube. Accordingly, because PTFE is a hydrophobic polymeric material, we conclude that hydrophobic interactions between PTFE and the organic groups on the QDs play a dominant role in the separation of the QDs from the perfusate samples. Optimization of Online In-Tube SPE for QDs. To optimize the analytical performance of the PTFE tube as a retention medium for the extraction of QDs from perfusate samples, we investigated the effects of three main parameters on the extraction efficiency: the pH of the sample, the extraction flow rate, and the dilution factor of sample. Effect of pH. To investigate the effect of pH on the retention of QDs on the inner wall of the PTFE tube, QD705-containing PBS solutions in the pH of 4-10 were transported through the PTFE tube at a flow rate of 50 µL min-1. Regardless of the acidity of the sample solution, because of the hydrophobic interactions Analytical Chemistry, Vol. 82, No. 17, September 1, 2010
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Figure 2. Signal intensity of QD705 plotted with respect to the flow rate during sample loading. All the data are normalized to the data of loading flow rate at 25 µL min-1.
between the QDs and PTFE, the extraction efficiency of the QDs remained consistent. To simplify the extraction process, we selected a pH of 7.4 (acidity of normal PBS solution) for our subsequent experiments. Effect of Sample Flow Rate. Because complete extraction of the QDs from the PPP samples would greatly improve the analytical performance, we studied the effect of the flow rate during sample loading. Figure 2 displays the variation in the final signal intensities at several different flow rates. We observed that the intensity of the signals obtained decreased upon increasing the flow rate, with the maximum retention of QDs occurring when the sample passed through the PTFE tube at 25 µL min-1. Because the mixed perfusate and buffer solution was transported directly to the PTFE tube at a flow rate of 20 µL min-1, satisfactory extraction of QDs was achieved during the sample loading step. Effect of Dilution Factor. After sampling from the extracellular space of rat liver using the PPP technique, the perfusates were very complicated mixtures, containing blood cells, proteins, and dissolved salts. To evaluate any possible negative effects of the extraction efficiency of QDs due to the involvement of blood matrix, we used various diluted blood samples, from 1:2 to 1:80, as sample matrixes. No apparent signal suppression occurred, even when 2-fold diluted blood was used as the sample matrix. Thus, we could adequately extract the QDs from the perfusate samples even when they comprised very complicated mixtures. Recovery of Push-Pull Probe. Next, we examined the practicability of the use of a microdialysis probe having a membrane with a molecular weight cutoff of 3 MDa to collect the QDs. We observed that only a very trivial amount of the QDs (
