Journal of Colloid and Interface Science 450 (2015) 388–395
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Journal of Colloid and Interface Science www.elsevier.com/locate/jcis
Adherence and interaction of cationic quantum dots on bacterial surfaces Cheng Yang a,b, Hao Xie a,b,⇑, Qi-Chang Li a,b, En-Jie Sun b, Bao-Lian Su a,c,⇑ a
State Key Laboratory of Advanced Technology for Materials Synthesis and Processing, Wuhan University of Technology, 122 Luoshi Road, Wuhan 430070, PR China School of Chemistry, Chemical Engineering and Life Sciences, Wuhan University of Technology, 122 Luoshi Road, Wuhan 430070, PR China c Laboratory of Inorganic Materials Chemistry, University of Namur, 61 rue de Bruxelles, Namur B-5000, Belgium b
g r a p h i c a l a b s t r a c t
a r t i c l e
i n f o
Article history: Received 3 February 2015 Accepted 23 March 2015 Available online 30 March 2015 Keywords: Quantum dots (QDs) Gram-positive bacteria Gram-negative bacteria Lipopolysaccharides (LPS) Bacterial surface Bacterial cell wall Teichoic-acids Fluorescence quenching Cell membrane Nanoparticles
a b s t r a c t Understanding molecular mechanisms of interactions between nanoparticles and bacteria is important and essential to develop nanotechnology for medical and environmental applications. Quantum dots (QDs) are specific nanoparticles with unique optical properties and high photochemical stability. In the present study, direct interactions were observed between cationic QDs and bacteria. Distinct fluorescence quenching patterns were developed when cationic QDs interacted with Gram negative and Gram positive bacteria. The aggregation of QDs on bacterial surface as well as fluorescence quenching depends upon the chemical composition and structure of the bacterial cell envelopes. The presence of lipopolysaccharide is unique to Gram-negative bacterial surface and provides negatively charge areas for absorbing cationic QDs. The effects of lipopolysaccharide were proved on fluorescence quenching of cationic QDs. In contrast to Gram-negative bacteria, the presence of teichoic acids is unique to Gram-positive bacterial cell wall and provides negatively charged sites for cationic QDs along the chain of teichoic-acid molecules, which may protect QDs from aggregation and fluorescence quenching. This study may not only provide insight into behaviors of QDs on bacterial cell surfaces but also open a new avenue for designing and applying QDs as biosensors in microbiology, medicine, and environmental science. Ó 2015 Elsevier Inc. All rights reserved.
⇑ Corresponding authors at: State Key Laboratory of Advanced Technology for Materials Synthesis and Processing, Wuhan University of Technology, 122 Luoshi Road, Wuhan 430070, PR China. E-mail addresses: [email protected] (H. Xie), [email protected] (B.-L. Su). http://dx.doi.org/10.1016/j.jcis.2015.03.041 0021-9797/Ó 2015 Elsevier Inc. All rights reserved.
C. Yang et al. / Journal of Colloid and Interface Science 450 (2015) 388–395
1. Introduction Nanotechnology has provided a basis for innovation in a wide range of researches and applications. Based on interactions between nanoparticles and cells or biomolecules, novel techniques have been developed and facilitated various biomedical applications including cancer detection [1], drug delivery [2], cell imaging [3], and proteins or bacteria identification [4,5]. Understanding molecular mechanisms of interactions between nanoparticles and cells or biomolecules facilitates developing novel nanotechnology, preventing environmental hazards, and reducing health risks of nanoparticles. In nature, cells are divided in two broad categories, eukaryotic cells and prokaryotic cells, depending on whether they contain nucleus. Internalization of nanoparticles into eukaryotic cells has been reported through endocytosis. Intracellular transport of nanoparticles has also been observed [6]. In contrast to eukaryotes, prokaryotes such as bacteria do not have an endocytosis system to acquire macromolecules. Nanoparticles may directly interact with and adhere or deposit on bacterial surface [7,8]. The multilayered bacteria cell envelope falls into one of two major groups [9,10]. Gram-negative bacteria are surrounded by two layers of membranes. The outer membrane mainly consists of proteins and lipopolysaccharide (LPS). The presence of LPS is unique to Gram-negative bacteria. Gram-positive bacteria lack an outer membrane but are surrounded by a thick layer of peptidoglycan which is threaded through with teichoic acids, a polymer being unique to Gram-positive bacterial cell wall [9]. When interact with the two types of bacterial surface, environmental substances including nanoparticles may exhibit distinct interaction patterns, which is essential to develop bacteria detectors or sensors [4], microorganisms hybrid devices [7], antibacterial nanoparticles [11], and evaluate environmental risks of nanoparticles [12]. Both types of bacterial surface contain many groups which can deprotonate to have negative charges, and adhere to positivelycharged materials [13,14]. Therefore, the electrostatic attraction and repulsion forces can make significant contributions to the interaction between cell surface and positively or negatively charged nanoparticles. Positively charged nanoparticles may possess an interaction mechanism different to negatively charged ones. Cationic nanoparticles such as CTAB-terminated nanospheres and cationic AuNPs can deposit or aggregate on bacterial surface [7,8]. The cationic, hydrophobic, monolayer- protected AuNPs may interact with bacteria through hydrophobic interaction with bacterial surface proteins. The bacterial surface structure is critical to the distinct aggregation patterns of AuNPs on Gram-positive and Gramnegative bacteria [8]. A recent report also showed that the negative or positive charge of nanoparticle would affect cell-membrane interactions [15]. Quantum dots (QDs) are specific nanoparticles with unique optical properties and high photochemical stability, and have already been widely used for cell imaging, molecular labeling and biosensing [16,17]. By taking QDs as representatives, interactions between nanoparticles and cell surface can be investigated with fluorescence techniques. Although scientists have observed interactions between negatively charged QDs and Gramnegative bacteria, there are few reports regarding interactions between positively charged QDs and bacteria [18]. Cysteamine-stabilized CdTe QDs are positively charged in neutral environment and have been reported linking to DNA through amino groups on the QDs surface [19]. In the present study, direct interactions were examined between cysteamine-stabilized CdTe/ ZnS core/shell QDs and bacteria. Distinct interaction profiles were observed between Gram-negative bacteria Escherichia coli and Gram-positive bacteria Bacillus subtilis. Roles of lipopolysaccharides were explored on fluorescence quenching of QDs. The present study focuses on behaviors and fluorescent characteristics of QDs
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when interact with bacteria, which is essential and important in designing and applying QDs as biosensors in microbiology, medicine, and environmental science. It is also helpful in understanding mechanisms of interactions between similar nanoparticles and bacteria. 2. Materials and methods 2.1. Synthesis of cysteamine-stabilized CdTe and CdTe/ZnS core/shell QDs Cysteamine-stabilized CdTe QDs were synthesized by a facile one-pot approach using sodium tellurite as the Te source. Typically, 545 mg of cysteamine hydrochloride and 365 mg of CdCl22.5H2O were added to 80 ml of deionized water. The pH was adjusted to 5.7–5.8 with NaOH. Then 17 mg of Na2TeO3 and 80 mg NaBH4 were added, followed by heating to 100 °C and refluxing until the photo-luminescence emission peak reached to 560 nm at the excitation wavelength 450 nm. For synthesis of CdTe/ZnS core/shell QDs, 29 mg of ZnSO47H2O and 272 mg of cysteamine hydrochloride were added to 40 ml of deionized water. The pH was adjusted to 5.7–5.8 with NaOH and mixed with 40 ml of freshly synthetized CdTe QDs solution, followed by adding 2 mg of Na2S9H2O. The mixtures were refluxed under 100 °C until the photo-luminescence emission peak reached to 570 nm at the excitation wavelength 450 nm. The QDs solution was stored at 4 °C for later use. Aliquots of the reaction mixture were removed at regular intervals for measuring UV absorption with a UV-2550 spectrophotometer (SHIMADZU) or analyzing photoluminescence with a LS55 fluorescence spectrometer (Perkin Elmer) at room temperature. 2.2. Bacterial strains and culturing Ten bacterial strains were investigated including E. coli DH5a, B. subtilis B168, Listeria monocytogenes CMCC(B) 54002, Pseudomonas aeruginosa ATCC 27853, Staphylococcus epidermidis CMCC(B) 269069, Proteus vulgaris CMCC(B) 49027, Salmonella paratyphi A CMCC(B) 50093, Staphyloccocus aureus, Bacillus cereus, Serratia sp. All bacteria were cultured in Luria–Bertani (LB) medium with shaking for four to five hours at 37 °C and harvested by centrifugation at 4000g for 10 min. 2.3. Cysteamine-stabilized QDs adsorption by bacteria Bacterial cells were washed and resuspended in 50 mM Tris–HCl, pH 7.0 (Buffer A). Typically, 100 ll of bacteria at specified cell density (OD600) were mixed with 200 ll of freshly prepared cysteamine-stabilized CdTe/ZnS core/shell QDs for 10 min. QDs-labeled bacterial cells were sedimented by centrifugation at 4000g for 10 min, followed by and washing with Buffer A, then resuspended in 300 ll of Buffer A for later use. 2.4. Fluorescence spectrometry The fluorescence spectra of QDs or QDs-labeled bacteria were recorded with a LS55 fluorescence spectrometer (Perkin-Elmer) at an excitation wave length of 450 nm. Bacterial cells labeled by QDs were separated by centrifugation. QDs-labeled bacterial cells and the supernatant were collected and analysed by fluorescence spectrometer, respectively. For monitoring the time-dependent fluorescence quench, 2.7 ml of bacterial cells at density of 0.3 OD600 were mixed with 200 ll of QDs and monitored immediately with a fluorescence spectrometer.
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2.5. Ethanol and trypsin treatment of bacteria Bacterial cells were either treated in 95% ethanol for 5 min or incubated in Buffer A containing 1% trypsin at 37 °C for 5 h, followed by sedimentation at 4000g for 10 min and washed twice with Buffer A, then resuspended in Buffer A for later use.
implying the incorporation of ZnS shell on the core CdTe QDs [22]. The size and concentration of as prepared CdTe/ZnS QDs were calculated as 3.1 nm and 4 106 mole/L by using an empirical formula [24]. Subsequent assays were performed with the CdTe/ZnS core/shell QDs. Typically, 200 ll of QDs were mixed with 100 ll of bacterial at a specified cell density for investigating interactions between QDs and bacteria.
2.6. Fluorescence quenching by LPS Lipopolysaccharides (LPS) from E. coli 055:B5 were purchased from Sigma. The LPS were dispersed in water. Different amount of the LPS were mixed with QDs and subjected for fluorescence spectrometry. For comparison, 100 ll of E. coli cells at various cell densities were mixed with QDs and subjected for fluorescence spectrometry. 2.7. Fluorescence microscopy Cell suspension was applied to a cover glass and analyzed with an inverted fluorescence microscope (IX71). Emissions were examined under UV-excitation. 2.8. Flow cytometric measurement Flow cytometric measurement was performed with a FACS Calibur (Becton Dickinson) flow cytometer. Samples were illuminated with an argon laser (488 nm), and the fluorescence was detected through pass filter FL2 channel. Sample voltages were set at 325 V for side scatter (SSC) and 670 V for detectors FL1. To eliminate influence of small non-bacterial particles, a threshold value of 58 was set on FSC (forward scatter). Cells were dispersed in Buffer A. About 20,000 cells were detected at a rate of approximately 1000–2000 events per second. 2.9. Transmission electron microscopy of bacteria Bacterial cells were resuspended in sterile water and loaded on a carbon-coated, 200 mesh copper grids, followed by examination with a transmission electron microscope (JEOL JEM 2000F). 2.10. Toxicity of QDs on bacterial growth Unlabeled or QDs-labeled bacterial cells were washed and resuspended in 300 ll of LB medium. An aliquot (150 ll) of bacteria was inoculated in 10 ml of LB medium, followed by shaking at 37 °C. Cell growth was monitored by measuring cell density at OD600. A set of serial dilutions of the bacterial cells is made and followed by plating on solidified LB medium. Cell viability was examined by counting the number of bacterial colonies developed on the plate and expressed as colony-forming units (CFU/ml). 3. Result 3.1. Synthesis and characterization of cysteamine-stabilized CdTe/ZnS core/shell QDs Aqueous phase synthesis of cysteamine-stabilized CdTe QDs was based on previous facile one-pot approaches [20,21]. To enhance the stability and reduce the toxicity, CdTe QDs were coated with ZnS by adding Zn2+ and S2 into the CdTe solution at 100 °C under reflux conditions [22,23]. The photo-luminescence emission and absorption spectra were compared between cysteamine-stabilized CdTe QDs and CdTe/ZnS core/shell QDs (Fig. 1). There was a red shift of the photo-luminescence emission peak and absorption peak between CdTe/ZnS QDs and CdTe QDs,
3.2. Direct interactions between cysteamine-stabilized QDs and bacteria E. coli and B. subtilis were used as representatives of Gram-negative and Gram-positive bacteria. Interactions between cysteaminestabilized QDs and bacteria were analyzed by means of fluorescence spectrophotometry, fluorescence microscopy, and cytometry. If not stated otherwise, the interaction time between QDs and bacteria is 10 min. Upon interaction with QDs, both E. coli and B. subtilis exhibited significant fluorescence increases, which were detected by fluorescence spectrophotometry and flow cytometry (Fig. 2, Panels A and B). Fluorescence microscopy also demonstrated that both bacteria were labeled with QDs (Fig. 2, Panels C, C0 , D, and D0 ). Complete adsorption of QDs by bacterial cells was observed by comparing the fluorescence spectrum of QDs solutions before and after interacting with bacteria (Fig. 2, Panel A). A red shift of the photo-luminescence emission of QDs was observed after interacting with bacteria, implying there was aggregation of QDs (Fig. 2, Panel A) [25,26]. When same amount of QDs interacted with a set of excessive amount of bacteria at cell densities ranging from 0.42 OD600 to 0.85 OD600, complete adsorption of QDs by E. coli or B. subtilis were confirmed by the observation that the fluorescence of QDs solution decreased to zero and fluorescence of bacteria significantly increased to a level. There were no significant fluorescence changes for each bacterium when different amount of cells were tested (Fig. 3, Panel A). Besides, there was significant fluorescence quenching of QDs absorbing by E. coli than that of B. subtilis (Fig. 3, Panel A). Therefore, it can be concluded, when QDs were absorbed by excessive amount of bacterial cells, the fluorescence and quenching patterns can be utilized to distinguish the two bacteria. The relationship was also investigated between the fluorescence change and interaction time of QDs with bacteria (Fig. 3, Panel B). E. coli exhibited a significantly faster decreasing rate of fluorescence than that of B. subtilis, implying the aggregation and precipitation of QDs is in progress. The toxicity of QDs to the two bacteria was tested and described as bacterial colony-forming units (CFU) before or after interaction with QDs (Fig. 3, Panel C), which suggested that B. subtilis was more susceptible to QDs than that of E. coli. 3.3. Effects of lipopolysaccharides on fluorescence quenching of QDs It is interesting that there was more fluorescence quenching of QDs when interacted with E. coli than B. subtilis. The major components of Gram-negative bacterial outer membrane are lipopolysaccharides (LPS) and lipoproteins which do not appear in Gram-positive bacteria [27]. LPS are large molecules consisting of a lipid and a polysaccharide with many phosphate groups. Approximately 75% of the E. coli cell surface is covered with LPS [27,28]. When QDs interacted with E. coli cells whose lipids including LPS on the cell surface were partially dissolved with ethanol, complete adsorption of QDs by bacterial cells was still observed, but the fluorescence quenching significantly reduced (Fig. 4, Panel A), suggesting that the lipids on cell surface made the major contribution to the fluorescence quenching. The roles of LPS in fluorescence quenching were supported by observations that there was only a slight fluorescence change when B. subtilis, a
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Fig. 1. A comparison of the emission spectra (Panel A) and corresponding absorption spectra (Panel B) between cysteamine-stabilized CdTe QDs and CdTe/ZnS core/shell QDs at the excitation wavelength of 450 nm.
Fig. 2. Interactions between QDs and bacteria examined by means of fluorescence-based technology. Panel A shows the fluorescence spectra of QDs-labeled bacteria and QDs solutions before and after interaction with bacteria. Panel B shows the flow cytometry spectra of bacteria before and after labeling with QDs. Panels C, C0 , D, and D0 show the bright-field microscopy (Panel C, D) and fluorescence microscopy images (Panels C0 , D0 ) of E. coli cells (Panels C, C0 ) and B. subtilis cells (Panels D, D0 ) after labeling with QDs.
Gram-positive bacterium with no LPS on its surface, was subjected to the same test (Fig. 4, Panel B). When QDs interacted with E. coli cells whose proteins on the cell surface were digested and removed by trypsin, complete adsorption of QDs by bacterial cells was observed and there was no significant change of the fluorescence emission spectrum between undigested and trypsin-digested cells (Fig. 4, Panel C), suggesting that surface proteins did not involve in
fluorescence quenching of QDs. It was also supported by observations that there were no significant fluorescence changes when trypsin-digested cells of B. subtilis were subjected to test (Fig. 4, Panel D). The interaction between QDs and pure LPS or E. coli cells was further investigated by titrating QDs solution with increasing amounts of LPS or E. coli (Fig. 4, Panels E and F). Results showed that both pure LPS and E. coli quenched the fluorescence of QDs
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Fig. 3. Panel A shows the QDs fluorescence intensity change when interacted with E. coli or B. subtilis cells at different cell densities. Panel B shows the relationship between the fluorescence intensity change and interaction time. Panel C shows bacterial colony-forming units (CFU) before or after interaction with QDs.
Fig. 4. The fluorescence emission spectra of QDs interacted with E. coli cells (Panel A) or B. subtilis cells (Panel B) treated by ethanol and E. coli cells (Panel C) or B. subtilis cells (panel D) treated by trypsin. The fluorescence emission spectra of QDs interacted with various amounts of E. coli cells (Panel E) and various amounts of LPS (Panel F).
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Fig. 5. TEM images of E. coli cells (Panels A, B, B0 ) and B. subtilis cells (Panels C, D, D0 ) after interacting with QDs. Panels B0 or D0 is corresponding to the higher-magnification view of the boxed area in Panels B or D.
in similar manners, implying there were specific interactions between LPS and QDs. Interaction details between QDs and E. coli cells were further examined by imaging QDs-labeled bacterial cells with transmission electron microscopy (Fig. 5). QDs aggregation was observed on QDs-labeled E. coli cell surface. However, there was no aggregation of QDs (Fig. 5, Panels D and D0 ) and significant fluorescence quenching (Fig. 2, Panel A) when QDs interacted with B. subtilis. 3.4. QDs-based rapid discrimination of Gram-positive and Gramnegative bacteria Since the interaction patterns of QDs to E. coli or B. subtilis are attributed to the bacterial surface characteristics, other Gram positive and Gram negative bacteria may interact with QDs in similar manners. In the present study, five representatives of each bacteria group were further investigated to verify this hypothesis (Fig. 6).
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Fig. 6. QDs-based discrimination of Gram-negative and Gram-positive bacteria.
Briefly, excessive living bacteria cells at high densities (OD600 > 2) were subjected to test to ensure the saturated adsorption of QDs and maximal fluorescence quenching. Same amount of QDs were mixed with bacteria for 10 min, followed by fluorescence measurements. Low fluorescence intensity was observed when QDs interacted with all Gram-negative bacteria. While, higher fluorescence intensity was observed when QDs interacted with all Gram-positive bacteria. The distinct interaction pattern of QDs to Gram-negative and Gram-positive bacteria (Fig. 6) suggests the potential of developing a simple method for rapid discrimination of the two types of bacteria.
4. Discussions Almost all biochemical reactions and physiological processes take place in aqueous environments. Good water dispersibility is essential for QDs in purposes of biological applications. Carboxyl quantum dots such as TGA, MPA, or MSA stabilized QDs are water dispersible and used in a wide variety of labeling and tracking applications [29–31]. However, it is not easy to employ these negatively charged carboxyl quantum dots in direct labeling of E. coli cells whose surface are also negatively charged [32]. The present study provides an alternative to label bacterial cells by using positively charged cysteamine-stabilized CdTe QDs based on direct interactions between bacteria and cysteamine-stabilized CdTe QDs (Figs. 2 and 7). The present study has observed that E. coli cells or pure LPS significantly quenched the fluorescence of cysteamine-stabilized CdTe QDs (Figs. 3, 4 and 7). The decrease of overall fluorescence intensity can be attributed to the QDs clustering in solutions or forming a solid nanocrystal film [25,26,33]. The fluorescence quenching reflects QDs clustering status on bacterial cell surface. QDs aggregation on E. coli cell surface is confirmed with transmission electron microscopy (Fig. 5). Considering the structural difference between Gram-negative and Gram-positive bacteria, B. subtilis lacks LPS
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Fig. 7. Illustration of interactions between QDs and bacteria.
which is the major component of the E. coli outer membrane. LPS are large molecules consisting of a lipid and a polysaccharide. Phosphate groups of LPS molecules or other phospholipids molecules on cell outer membrane form negatively charged area on E. coli cell surface (Fig. 7), on which positively charged QDs can be absorbed and aggregated. This has been confirmed by the observation that fluorescence quenching occurred when cysteamine-stabilized CdTe QDs interacted with pure LPS, which would form micelles and lead to QDs aggregation [34,35]. When E. coli interacted with cysteamine-stabilized CdTe QDs, the fluorescence quenching of QDs was related to time (Fig. 3, Panel B), implying the movement and accumulation of LPS-adhered QDs on cell surface. In contrast to Gram-negative bacteria, the surface of Grampositive bacteria such as B. subtilis is mainly composed of proteins, teichoic-acid and peptidoglycan. Ionizable functional groups are normally associated with peptidoglycan and polymers such as teichoic-acids. Teichoic-acids including teichoic and teichuronic acids are long chain molecules which are composed of negatively charged
repeating units. The length of teichoic-acids exposing to the cell surface is more than 40 nm [36]. The negatively charged groups are along the long chain of teichoic-acid molecules on B. subtilis cell surface and provide a static distribution of binding sites for positively charged QDs (Fig. 7). Upon interacting with B. subtilis, QDs will bind to these binding sites and not aggregate. Therefore, no significant fluorescence quenching occurred when B. subtilis was labeled with positively charged QDs (Fig. 2, Panel A; Fig. 3, Panels A and B). Comparing with Gram-positive bacteria, there is an extra lipid layer surrounding the Gram-negative bacterial cells. When exposed to environmental agents such as QDs, the extra lipid layer may provide Gram-negative bacteria with more protections than Gram-positive bacteria. This explains that B. subtilis was more susceptible to QDs than that of E. coli (Fig. 3, Panel C). When the lipid membrane on E. coli cell surface were partially dissolved in ethanol, there were still many other negatively charged groups distributing on the peptidoglycan layer of E. coli cells [27,28]. This explains that QDs could still be completely absorbed by E. coli cells treating with ethanol (Fig. 4, Panel A).
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A recent study showed that aggregations of 6 nm AuNPs on bacterial surface through hydrophobic interactions between proteins and AuNPs. And aggregation of 2 nm AuNPs on E. coli outer membrane were time-dependent and led to nanoparticle towers by resembling arrested precursors to the outer membrane vesicles [8], suggesting direct interactions between AuNPs and the cell surface. In the present study, no significant fluorescence change was observed after the surface proteins were removed (Fig. 4, Panels C and D), implying that proteins on cell surface did not involve in QDs/bacteria interactions. Actually, it was phospholipids molecules such as LPS on the membrane were demonstrated to involve in the adsorption and aggregation of QDs on E. coli cells. Since QDs used in the present study were modified with hydrophilic surface, it is not likely that hydrophobic interaction led to QDs/bacteria interactions. Therefore, it can be electrostatic attraction makes major contributions to QDs/bacteria interactions. The observation of time-dependent QDs aggregation on E. coli outer membrane (Fig. 3, Panel B, and Fig. 5) suggested that QDs may interact with E. coli cell surface in a manner similar to that of 2 nm AuNPs. In addition to LPS, positively charged QDs may also interact with other negatively charged molecules on bacterial surface, such as negatively charged phospholipid groups on bacterial inner membranes or eukaryotic cell membranes. Aggregation and fluorescence quenching of QDs may also occur upon interaction with these surface. Actually, significant fluorescence quenching did occur when QDs interacted with eukaryotic cells of Tetrahymena thermophile or ‘‘naked’’ cells of E. coli or B. subtilis whose cell wall were removed by lysozyme to expose their inner membrane (for details, see the Supporting Information). Therefore, positively charged QDs can be broadly used in interaction with both bacterial and eukaryotic cells. 5. Conclusions The present study reports the behavior and direct interactions between the cysteamine-stabilized CdTe QDs and bacterial cells. Distinct fluorescence quenching patterns were developed when Gram negative or Gram positive bacteria cationic interacted with QDs. Aggregation of QDs and fluorescence quenching was observed and related to chemical compositions and structures of bacterial cell envelopes. Negatively charged groups of molecules such as LPS may also play an important role in the interaction between hydrophilic cationic nanoparticles and bacterial cells. The potential application was also demonstrated in rapid discrimination of Gram-negative and Gram-positive bacteria. The present study may not only provide insight into behaviors of QDs on bacterial cell surfaces but also open a new avenue for designing and applying QDs as biosensors in microbiology, medicine, and environmental science. Further fluorescence-based detections and measurements of intact bacterial cells can be achieved based on the direct interactions between cationic QDs and bacterial cell surface. Acknowledgments This work is financially supported by ‘‘the Fundamental Research Funds for the Central Universities’’ (WHUT, 2013-Ia037). B.-L. Su also acknowledges the Chinese Central Government for an ‘‘Expert of the State’’ position in the program of ‘‘Thousand
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talents’’, the Chinese Ministry of Education for the program for Changjiang Scholars and Innovative Research Team (IRT1169) at the Wuhan University of Technology. Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jcis.2015.03.041. References [1] X. Huang, I.H. El-Sayed, W. Qian, M.A. El-Sayed, J. Am. Chem. Soc. 128 (2006) 2115–2120. [2] J. Panyam, V. Labhasetwar, Adv. Drug Deliv. Rev. 64 (2012) 61–71. [3] L. Zhu, W. Wu, M.Q. Zhu, J.J. Han, J.K. Hurst, A.D.Q. Li, J. Am. Chem. Soc. 129 (2007) 3524–3526. [4] R.L. Phillips, O.R. Miranda, C.C. You, V.M. Rotello, U.H.F. Bunz, Angew. Chem. Int. Ed. 47 (2008) 2590–2594. [5] C.C. You, O.R. Miranda, B. Gider, P.S. Ghosh, I.B. Kim, B. Erdogan, S.A. Krovi, U.H.F. Bunz, V.M. Rotello, Nat. Nanotechnol. 2 (2007) 318–323. [6] T.G. Iversen, T. Skotland, K. Sandvig, Nano Today 6 (2011) 176–185. [7] V. Berry, A. Gole, S. Kundu, C.J. Murphy, R.F. Saraf, J. Am. Chem. Soc. 127 (2005) 17600–17601. [8] S.C. Hayden, G. Zhao, K. Saha, R.L. Phillips, X. Li, O.R. Miranda, V.M. Rotello, M.A. El-Sayed, I.S. Krey, U.H.F. Bunz, J. Am. Chem. Soc. 134 (2012) 6920–6923. [9] T.J. Silhavy, D. Kahne, S. Walker, Cold Spring Harb. Perspect Biol. 2 (2010) a000414. [10] H.R. Perkins, Bacteriol. Rev. 27 (1963) 18–55. [11] M.J. Hajipour, K.M. Fromm, A.A. Ashkarran, D.J. Aberasturi, I.R. Larramendi, T. Rojo, V. Serpooshan, W.J. Parak, M. Mahmoudi, Trends. Biotechnol. 30 (2012) 499–511. [12] M.A. Maurer-Jones, I.L. Gunsolus, C.J. Murphy, C.L. Haynes, Anal. Chem. 85 (2013) 3036–3049. [13] M.M. Sharma, Y.I. Chang, T.F. Yen, Colloid. Surface. 16 (1985) 193–206. [14] N. Yee, D.A. Fowle, F.G. Ferris, Geochim. Cosmochim. Ac. 68 (2004) 3657–3664. [15] S. Tatur, M. Maccarini, R. Barker, A. Nelson, G. Fragneto, Langmuir 29 (2013) 6606–6614. [16] M.M. Barroso, J. Histochem. Cytochem. 59 (2011) 237–251. [17] X. Michalet, F.F. Pinaud, L.A. Bentolila, J.M. Tsay, S. Doose, J.J. Li, G. Sundaresan, A.M. Wu, S.S. Gambhir, S. Weiss, Science 307 (2005) 538–544. [18] L. Lai, C. Lin, C.Q. Xiao, Z.Q. Xu, X.L. Han, L. Fu, D.W. Li, P. Mei, F.L. Jiang, Q.L. Guo, Y. Liu, J. Colloid. Interf. Sci. 389 (2013) 61–70. [19] Q. Chen, D. Yang, S. Xu, Anal. Lett. 41 (2008) 1964–1974. [20] R. Wang, Y. Wang, Q. Feng, L. Zhou, F. Gong, Y. Lan, Mater. Lett. 66 (2012) 261– 263. [21] Q.F. Chen, W.X. Wang, Y.X. Ge, M.Y. Li, S.K. Xu, X.J. Zhang, Chin. J. Anal. Chem. 35 (2007) 135–138. [22] Y. He, H.T. Lu, L.M. Sai, Y.Y. Su, M. Hu, C.H. Fan, W. Huang, L.H. Wang, Adv. Mater. 20 (2008) 3416–3421. [23] F.D. Wei, Y. Li n, Y.Z. Wu, X. Sun, L.P. Liu, P. Zhou, Q. Hu, Anal. Methods 6 (2014) 482–489. [24] W.W. Yu, L. Qu, W. Guo, X. Peng, Chem. Mater. 15 (2003) 2854–2860. [25] R. Koole, P. Liljeroth, C.M. Donega, D.L. Vanmaekelbergh, A. Meijerink, J. Am. Chem. Soc 128 (2006) 10436–10441. [26] S. Mayilo, J. Hilhorst, A.S. Susha, C. Hohl, T. Franzl, T.A. Klar, A.L. Rogach, J. Feldmann, J. Phys. Chem. C 112 (2008) 14589–14594. [27] H.L. Alakomi, Weakening of the Gram-Negative Bacterial Outer Membrane, a Tool for Increasing Microbiological Safety, VTT Publications, Espoo, 2007. [28] M.P. Bos, J. Tommassen, Curr. Opin. Microbiol. 7 (2004) 610–616. [29] M. Hu, J. Yan, Y. He, H. Lu, L. Weng, S. Song, C. Fan, L. Wang, ACS Nano 4 (2010) 488–494. [30] Y. Williams, A. Sukhanova, M. Nowostawska, A.M. Davies, S. Mitchell, V. Oleinikov, Y. Gun’ko, I. Nabiev, D. Kelleher, Y. Volkov, Small 5 (2009) 2581– 2588. [31] M.A. Walling, J.A. Novak, J.R.E. Shepard, Int. J. Mol. Sci. 10 (2009) 441–491. [32] C. Yang, H. Xie, Y. Li, J.K. Zhang, B.L. Su, J. Colloid. Interf. Sci. 396 (2013) 438– 444. [33] A.S. Susha, A.M. Javier, W.J. Parak, A.L. Rogach, Colloid. Surf. A-Physicochem. Eng. Asp. 281 (2006) 40–43. [34] L. Leive, D.C. Morrison, Methods. Enzymol. 28 (1972) 254–262. [35] B. Jann, K. Reske, K. Jann, Eur. J. Biochem. 60 (1975) 239–246. [36] D.C. Birdsell, R.J. Doyle, M. Morgenstern, J. Bacteriol. 121 (1975) 726–734.
Contents lists available at ScienceDirect
Journal of Colloid and Interface Science www.elsevier.com/locate/jcis
Adherence and interaction of cationic quantum dots on bacterial surfaces Cheng Yang a,b, Hao Xie a,b,⇑, Qi-Chang Li a,b, En-Jie Sun b, Bao-Lian Su a,c,⇑ a
State Key Laboratory of Advanced Technology for Materials Synthesis and Processing, Wuhan University of Technology, 122 Luoshi Road, Wuhan 430070, PR China School of Chemistry, Chemical Engineering and Life Sciences, Wuhan University of Technology, 122 Luoshi Road, Wuhan 430070, PR China c Laboratory of Inorganic Materials Chemistry, University of Namur, 61 rue de Bruxelles, Namur B-5000, Belgium b
g r a p h i c a l a b s t r a c t
a r t i c l e
i n f o
Article history: Received 3 February 2015 Accepted 23 March 2015 Available online 30 March 2015 Keywords: Quantum dots (QDs) Gram-positive bacteria Gram-negative bacteria Lipopolysaccharides (LPS) Bacterial surface Bacterial cell wall Teichoic-acids Fluorescence quenching Cell membrane Nanoparticles
a b s t r a c t Understanding molecular mechanisms of interactions between nanoparticles and bacteria is important and essential to develop nanotechnology for medical and environmental applications. Quantum dots (QDs) are specific nanoparticles with unique optical properties and high photochemical stability. In the present study, direct interactions were observed between cationic QDs and bacteria. Distinct fluorescence quenching patterns were developed when cationic QDs interacted with Gram negative and Gram positive bacteria. The aggregation of QDs on bacterial surface as well as fluorescence quenching depends upon the chemical composition and structure of the bacterial cell envelopes. The presence of lipopolysaccharide is unique to Gram-negative bacterial surface and provides negatively charge areas for absorbing cationic QDs. The effects of lipopolysaccharide were proved on fluorescence quenching of cationic QDs. In contrast to Gram-negative bacteria, the presence of teichoic acids is unique to Gram-positive bacterial cell wall and provides negatively charged sites for cationic QDs along the chain of teichoic-acid molecules, which may protect QDs from aggregation and fluorescence quenching. This study may not only provide insight into behaviors of QDs on bacterial cell surfaces but also open a new avenue for designing and applying QDs as biosensors in microbiology, medicine, and environmental science. Ó 2015 Elsevier Inc. All rights reserved.
⇑ Corresponding authors at: State Key Laboratory of Advanced Technology for Materials Synthesis and Processing, Wuhan University of Technology, 122 Luoshi Road, Wuhan 430070, PR China. E-mail addresses: [email protected] (H. Xie), [email protected] (B.-L. Su). http://dx.doi.org/10.1016/j.jcis.2015.03.041 0021-9797/Ó 2015 Elsevier Inc. All rights reserved.
C. Yang et al. / Journal of Colloid and Interface Science 450 (2015) 388–395
1. Introduction Nanotechnology has provided a basis for innovation in a wide range of researches and applications. Based on interactions between nanoparticles and cells or biomolecules, novel techniques have been developed and facilitated various biomedical applications including cancer detection [1], drug delivery [2], cell imaging [3], and proteins or bacteria identification [4,5]. Understanding molecular mechanisms of interactions between nanoparticles and cells or biomolecules facilitates developing novel nanotechnology, preventing environmental hazards, and reducing health risks of nanoparticles. In nature, cells are divided in two broad categories, eukaryotic cells and prokaryotic cells, depending on whether they contain nucleus. Internalization of nanoparticles into eukaryotic cells has been reported through endocytosis. Intracellular transport of nanoparticles has also been observed [6]. In contrast to eukaryotes, prokaryotes such as bacteria do not have an endocytosis system to acquire macromolecules. Nanoparticles may directly interact with and adhere or deposit on bacterial surface [7,8]. The multilayered bacteria cell envelope falls into one of two major groups [9,10]. Gram-negative bacteria are surrounded by two layers of membranes. The outer membrane mainly consists of proteins and lipopolysaccharide (LPS). The presence of LPS is unique to Gram-negative bacteria. Gram-positive bacteria lack an outer membrane but are surrounded by a thick layer of peptidoglycan which is threaded through with teichoic acids, a polymer being unique to Gram-positive bacterial cell wall [9]. When interact with the two types of bacterial surface, environmental substances including nanoparticles may exhibit distinct interaction patterns, which is essential to develop bacteria detectors or sensors [4], microorganisms hybrid devices [7], antibacterial nanoparticles [11], and evaluate environmental risks of nanoparticles [12]. Both types of bacterial surface contain many groups which can deprotonate to have negative charges, and adhere to positivelycharged materials [13,14]. Therefore, the electrostatic attraction and repulsion forces can make significant contributions to the interaction between cell surface and positively or negatively charged nanoparticles. Positively charged nanoparticles may possess an interaction mechanism different to negatively charged ones. Cationic nanoparticles such as CTAB-terminated nanospheres and cationic AuNPs can deposit or aggregate on bacterial surface [7,8]. The cationic, hydrophobic, monolayer- protected AuNPs may interact with bacteria through hydrophobic interaction with bacterial surface proteins. The bacterial surface structure is critical to the distinct aggregation patterns of AuNPs on Gram-positive and Gramnegative bacteria [8]. A recent report also showed that the negative or positive charge of nanoparticle would affect cell-membrane interactions [15]. Quantum dots (QDs) are specific nanoparticles with unique optical properties and high photochemical stability, and have already been widely used for cell imaging, molecular labeling and biosensing [16,17]. By taking QDs as representatives, interactions between nanoparticles and cell surface can be investigated with fluorescence techniques. Although scientists have observed interactions between negatively charged QDs and Gramnegative bacteria, there are few reports regarding interactions between positively charged QDs and bacteria [18]. Cysteamine-stabilized CdTe QDs are positively charged in neutral environment and have been reported linking to DNA through amino groups on the QDs surface [19]. In the present study, direct interactions were examined between cysteamine-stabilized CdTe/ ZnS core/shell QDs and bacteria. Distinct interaction profiles were observed between Gram-negative bacteria Escherichia coli and Gram-positive bacteria Bacillus subtilis. Roles of lipopolysaccharides were explored on fluorescence quenching of QDs. The present study focuses on behaviors and fluorescent characteristics of QDs
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when interact with bacteria, which is essential and important in designing and applying QDs as biosensors in microbiology, medicine, and environmental science. It is also helpful in understanding mechanisms of interactions between similar nanoparticles and bacteria. 2. Materials and methods 2.1. Synthesis of cysteamine-stabilized CdTe and CdTe/ZnS core/shell QDs Cysteamine-stabilized CdTe QDs were synthesized by a facile one-pot approach using sodium tellurite as the Te source. Typically, 545 mg of cysteamine hydrochloride and 365 mg of CdCl22.5H2O were added to 80 ml of deionized water. The pH was adjusted to 5.7–5.8 with NaOH. Then 17 mg of Na2TeO3 and 80 mg NaBH4 were added, followed by heating to 100 °C and refluxing until the photo-luminescence emission peak reached to 560 nm at the excitation wavelength 450 nm. For synthesis of CdTe/ZnS core/shell QDs, 29 mg of ZnSO47H2O and 272 mg of cysteamine hydrochloride were added to 40 ml of deionized water. The pH was adjusted to 5.7–5.8 with NaOH and mixed with 40 ml of freshly synthetized CdTe QDs solution, followed by adding 2 mg of Na2S9H2O. The mixtures were refluxed under 100 °C until the photo-luminescence emission peak reached to 570 nm at the excitation wavelength 450 nm. The QDs solution was stored at 4 °C for later use. Aliquots of the reaction mixture were removed at regular intervals for measuring UV absorption with a UV-2550 spectrophotometer (SHIMADZU) or analyzing photoluminescence with a LS55 fluorescence spectrometer (Perkin Elmer) at room temperature. 2.2. Bacterial strains and culturing Ten bacterial strains were investigated including E. coli DH5a, B. subtilis B168, Listeria monocytogenes CMCC(B) 54002, Pseudomonas aeruginosa ATCC 27853, Staphylococcus epidermidis CMCC(B) 269069, Proteus vulgaris CMCC(B) 49027, Salmonella paratyphi A CMCC(B) 50093, Staphyloccocus aureus, Bacillus cereus, Serratia sp. All bacteria were cultured in Luria–Bertani (LB) medium with shaking for four to five hours at 37 °C and harvested by centrifugation at 4000g for 10 min. 2.3. Cysteamine-stabilized QDs adsorption by bacteria Bacterial cells were washed and resuspended in 50 mM Tris–HCl, pH 7.0 (Buffer A). Typically, 100 ll of bacteria at specified cell density (OD600) were mixed with 200 ll of freshly prepared cysteamine-stabilized CdTe/ZnS core/shell QDs for 10 min. QDs-labeled bacterial cells were sedimented by centrifugation at 4000g for 10 min, followed by and washing with Buffer A, then resuspended in 300 ll of Buffer A for later use. 2.4. Fluorescence spectrometry The fluorescence spectra of QDs or QDs-labeled bacteria were recorded with a LS55 fluorescence spectrometer (Perkin-Elmer) at an excitation wave length of 450 nm. Bacterial cells labeled by QDs were separated by centrifugation. QDs-labeled bacterial cells and the supernatant were collected and analysed by fluorescence spectrometer, respectively. For monitoring the time-dependent fluorescence quench, 2.7 ml of bacterial cells at density of 0.3 OD600 were mixed with 200 ll of QDs and monitored immediately with a fluorescence spectrometer.
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2.5. Ethanol and trypsin treatment of bacteria Bacterial cells were either treated in 95% ethanol for 5 min or incubated in Buffer A containing 1% trypsin at 37 °C for 5 h, followed by sedimentation at 4000g for 10 min and washed twice with Buffer A, then resuspended in Buffer A for later use.
implying the incorporation of ZnS shell on the core CdTe QDs [22]. The size and concentration of as prepared CdTe/ZnS QDs were calculated as 3.1 nm and 4 106 mole/L by using an empirical formula [24]. Subsequent assays were performed with the CdTe/ZnS core/shell QDs. Typically, 200 ll of QDs were mixed with 100 ll of bacterial at a specified cell density for investigating interactions between QDs and bacteria.
2.6. Fluorescence quenching by LPS Lipopolysaccharides (LPS) from E. coli 055:B5 were purchased from Sigma. The LPS were dispersed in water. Different amount of the LPS were mixed with QDs and subjected for fluorescence spectrometry. For comparison, 100 ll of E. coli cells at various cell densities were mixed with QDs and subjected for fluorescence spectrometry. 2.7. Fluorescence microscopy Cell suspension was applied to a cover glass and analyzed with an inverted fluorescence microscope (IX71). Emissions were examined under UV-excitation. 2.8. Flow cytometric measurement Flow cytometric measurement was performed with a FACS Calibur (Becton Dickinson) flow cytometer. Samples were illuminated with an argon laser (488 nm), and the fluorescence was detected through pass filter FL2 channel. Sample voltages were set at 325 V for side scatter (SSC) and 670 V for detectors FL1. To eliminate influence of small non-bacterial particles, a threshold value of 58 was set on FSC (forward scatter). Cells were dispersed in Buffer A. About 20,000 cells were detected at a rate of approximately 1000–2000 events per second. 2.9. Transmission electron microscopy of bacteria Bacterial cells were resuspended in sterile water and loaded on a carbon-coated, 200 mesh copper grids, followed by examination with a transmission electron microscope (JEOL JEM 2000F). 2.10. Toxicity of QDs on bacterial growth Unlabeled or QDs-labeled bacterial cells were washed and resuspended in 300 ll of LB medium. An aliquot (150 ll) of bacteria was inoculated in 10 ml of LB medium, followed by shaking at 37 °C. Cell growth was monitored by measuring cell density at OD600. A set of serial dilutions of the bacterial cells is made and followed by plating on solidified LB medium. Cell viability was examined by counting the number of bacterial colonies developed on the plate and expressed as colony-forming units (CFU/ml). 3. Result 3.1. Synthesis and characterization of cysteamine-stabilized CdTe/ZnS core/shell QDs Aqueous phase synthesis of cysteamine-stabilized CdTe QDs was based on previous facile one-pot approaches [20,21]. To enhance the stability and reduce the toxicity, CdTe QDs were coated with ZnS by adding Zn2+ and S2 into the CdTe solution at 100 °C under reflux conditions [22,23]. The photo-luminescence emission and absorption spectra were compared between cysteamine-stabilized CdTe QDs and CdTe/ZnS core/shell QDs (Fig. 1). There was a red shift of the photo-luminescence emission peak and absorption peak between CdTe/ZnS QDs and CdTe QDs,
3.2. Direct interactions between cysteamine-stabilized QDs and bacteria E. coli and B. subtilis were used as representatives of Gram-negative and Gram-positive bacteria. Interactions between cysteaminestabilized QDs and bacteria were analyzed by means of fluorescence spectrophotometry, fluorescence microscopy, and cytometry. If not stated otherwise, the interaction time between QDs and bacteria is 10 min. Upon interaction with QDs, both E. coli and B. subtilis exhibited significant fluorescence increases, which were detected by fluorescence spectrophotometry and flow cytometry (Fig. 2, Panels A and B). Fluorescence microscopy also demonstrated that both bacteria were labeled with QDs (Fig. 2, Panels C, C0 , D, and D0 ). Complete adsorption of QDs by bacterial cells was observed by comparing the fluorescence spectrum of QDs solutions before and after interacting with bacteria (Fig. 2, Panel A). A red shift of the photo-luminescence emission of QDs was observed after interacting with bacteria, implying there was aggregation of QDs (Fig. 2, Panel A) [25,26]. When same amount of QDs interacted with a set of excessive amount of bacteria at cell densities ranging from 0.42 OD600 to 0.85 OD600, complete adsorption of QDs by E. coli or B. subtilis were confirmed by the observation that the fluorescence of QDs solution decreased to zero and fluorescence of bacteria significantly increased to a level. There were no significant fluorescence changes for each bacterium when different amount of cells were tested (Fig. 3, Panel A). Besides, there was significant fluorescence quenching of QDs absorbing by E. coli than that of B. subtilis (Fig. 3, Panel A). Therefore, it can be concluded, when QDs were absorbed by excessive amount of bacterial cells, the fluorescence and quenching patterns can be utilized to distinguish the two bacteria. The relationship was also investigated between the fluorescence change and interaction time of QDs with bacteria (Fig. 3, Panel B). E. coli exhibited a significantly faster decreasing rate of fluorescence than that of B. subtilis, implying the aggregation and precipitation of QDs is in progress. The toxicity of QDs to the two bacteria was tested and described as bacterial colony-forming units (CFU) before or after interaction with QDs (Fig. 3, Panel C), which suggested that B. subtilis was more susceptible to QDs than that of E. coli. 3.3. Effects of lipopolysaccharides on fluorescence quenching of QDs It is interesting that there was more fluorescence quenching of QDs when interacted with E. coli than B. subtilis. The major components of Gram-negative bacterial outer membrane are lipopolysaccharides (LPS) and lipoproteins which do not appear in Gram-positive bacteria [27]. LPS are large molecules consisting of a lipid and a polysaccharide with many phosphate groups. Approximately 75% of the E. coli cell surface is covered with LPS [27,28]. When QDs interacted with E. coli cells whose lipids including LPS on the cell surface were partially dissolved with ethanol, complete adsorption of QDs by bacterial cells was still observed, but the fluorescence quenching significantly reduced (Fig. 4, Panel A), suggesting that the lipids on cell surface made the major contribution to the fluorescence quenching. The roles of LPS in fluorescence quenching were supported by observations that there was only a slight fluorescence change when B. subtilis, a
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Fig. 1. A comparison of the emission spectra (Panel A) and corresponding absorption spectra (Panel B) between cysteamine-stabilized CdTe QDs and CdTe/ZnS core/shell QDs at the excitation wavelength of 450 nm.
Fig. 2. Interactions between QDs and bacteria examined by means of fluorescence-based technology. Panel A shows the fluorescence spectra of QDs-labeled bacteria and QDs solutions before and after interaction with bacteria. Panel B shows the flow cytometry spectra of bacteria before and after labeling with QDs. Panels C, C0 , D, and D0 show the bright-field microscopy (Panel C, D) and fluorescence microscopy images (Panels C0 , D0 ) of E. coli cells (Panels C, C0 ) and B. subtilis cells (Panels D, D0 ) after labeling with QDs.
Gram-positive bacterium with no LPS on its surface, was subjected to the same test (Fig. 4, Panel B). When QDs interacted with E. coli cells whose proteins on the cell surface were digested and removed by trypsin, complete adsorption of QDs by bacterial cells was observed and there was no significant change of the fluorescence emission spectrum between undigested and trypsin-digested cells (Fig. 4, Panel C), suggesting that surface proteins did not involve in
fluorescence quenching of QDs. It was also supported by observations that there were no significant fluorescence changes when trypsin-digested cells of B. subtilis were subjected to test (Fig. 4, Panel D). The interaction between QDs and pure LPS or E. coli cells was further investigated by titrating QDs solution with increasing amounts of LPS or E. coli (Fig. 4, Panels E and F). Results showed that both pure LPS and E. coli quenched the fluorescence of QDs
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Fig. 3. Panel A shows the QDs fluorescence intensity change when interacted with E. coli or B. subtilis cells at different cell densities. Panel B shows the relationship between the fluorescence intensity change and interaction time. Panel C shows bacterial colony-forming units (CFU) before or after interaction with QDs.
Fig. 4. The fluorescence emission spectra of QDs interacted with E. coli cells (Panel A) or B. subtilis cells (Panel B) treated by ethanol and E. coli cells (Panel C) or B. subtilis cells (panel D) treated by trypsin. The fluorescence emission spectra of QDs interacted with various amounts of E. coli cells (Panel E) and various amounts of LPS (Panel F).
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Fig. 5. TEM images of E. coli cells (Panels A, B, B0 ) and B. subtilis cells (Panels C, D, D0 ) after interacting with QDs. Panels B0 or D0 is corresponding to the higher-magnification view of the boxed area in Panels B or D.
in similar manners, implying there were specific interactions between LPS and QDs. Interaction details between QDs and E. coli cells were further examined by imaging QDs-labeled bacterial cells with transmission electron microscopy (Fig. 5). QDs aggregation was observed on QDs-labeled E. coli cell surface. However, there was no aggregation of QDs (Fig. 5, Panels D and D0 ) and significant fluorescence quenching (Fig. 2, Panel A) when QDs interacted with B. subtilis. 3.4. QDs-based rapid discrimination of Gram-positive and Gramnegative bacteria Since the interaction patterns of QDs to E. coli or B. subtilis are attributed to the bacterial surface characteristics, other Gram positive and Gram negative bacteria may interact with QDs in similar manners. In the present study, five representatives of each bacteria group were further investigated to verify this hypothesis (Fig. 6).
Fluorescence intensity (a.u.)
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Gram negative Gram positive
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sp P. vu . lg ar is S. ty E. ph c im oli ur iu m S. au r B. eus su bt ili L. s m B. on ce oc re yt us og S. en ep es id er m id is
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Fig. 6. QDs-based discrimination of Gram-negative and Gram-positive bacteria.
Briefly, excessive living bacteria cells at high densities (OD600 > 2) were subjected to test to ensure the saturated adsorption of QDs and maximal fluorescence quenching. Same amount of QDs were mixed with bacteria for 10 min, followed by fluorescence measurements. Low fluorescence intensity was observed when QDs interacted with all Gram-negative bacteria. While, higher fluorescence intensity was observed when QDs interacted with all Gram-positive bacteria. The distinct interaction pattern of QDs to Gram-negative and Gram-positive bacteria (Fig. 6) suggests the potential of developing a simple method for rapid discrimination of the two types of bacteria.
4. Discussions Almost all biochemical reactions and physiological processes take place in aqueous environments. Good water dispersibility is essential for QDs in purposes of biological applications. Carboxyl quantum dots such as TGA, MPA, or MSA stabilized QDs are water dispersible and used in a wide variety of labeling and tracking applications [29–31]. However, it is not easy to employ these negatively charged carboxyl quantum dots in direct labeling of E. coli cells whose surface are also negatively charged [32]. The present study provides an alternative to label bacterial cells by using positively charged cysteamine-stabilized CdTe QDs based on direct interactions between bacteria and cysteamine-stabilized CdTe QDs (Figs. 2 and 7). The present study has observed that E. coli cells or pure LPS significantly quenched the fluorescence of cysteamine-stabilized CdTe QDs (Figs. 3, 4 and 7). The decrease of overall fluorescence intensity can be attributed to the QDs clustering in solutions or forming a solid nanocrystal film [25,26,33]. The fluorescence quenching reflects QDs clustering status on bacterial cell surface. QDs aggregation on E. coli cell surface is confirmed with transmission electron microscopy (Fig. 5). Considering the structural difference between Gram-negative and Gram-positive bacteria, B. subtilis lacks LPS
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Fig. 7. Illustration of interactions between QDs and bacteria.
which is the major component of the E. coli outer membrane. LPS are large molecules consisting of a lipid and a polysaccharide. Phosphate groups of LPS molecules or other phospholipids molecules on cell outer membrane form negatively charged area on E. coli cell surface (Fig. 7), on which positively charged QDs can be absorbed and aggregated. This has been confirmed by the observation that fluorescence quenching occurred when cysteamine-stabilized CdTe QDs interacted with pure LPS, which would form micelles and lead to QDs aggregation [34,35]. When E. coli interacted with cysteamine-stabilized CdTe QDs, the fluorescence quenching of QDs was related to time (Fig. 3, Panel B), implying the movement and accumulation of LPS-adhered QDs on cell surface. In contrast to Gram-negative bacteria, the surface of Grampositive bacteria such as B. subtilis is mainly composed of proteins, teichoic-acid and peptidoglycan. Ionizable functional groups are normally associated with peptidoglycan and polymers such as teichoic-acids. Teichoic-acids including teichoic and teichuronic acids are long chain molecules which are composed of negatively charged
repeating units. The length of teichoic-acids exposing to the cell surface is more than 40 nm [36]. The negatively charged groups are along the long chain of teichoic-acid molecules on B. subtilis cell surface and provide a static distribution of binding sites for positively charged QDs (Fig. 7). Upon interacting with B. subtilis, QDs will bind to these binding sites and not aggregate. Therefore, no significant fluorescence quenching occurred when B. subtilis was labeled with positively charged QDs (Fig. 2, Panel A; Fig. 3, Panels A and B). Comparing with Gram-positive bacteria, there is an extra lipid layer surrounding the Gram-negative bacterial cells. When exposed to environmental agents such as QDs, the extra lipid layer may provide Gram-negative bacteria with more protections than Gram-positive bacteria. This explains that B. subtilis was more susceptible to QDs than that of E. coli (Fig. 3, Panel C). When the lipid membrane on E. coli cell surface were partially dissolved in ethanol, there were still many other negatively charged groups distributing on the peptidoglycan layer of E. coli cells [27,28]. This explains that QDs could still be completely absorbed by E. coli cells treating with ethanol (Fig. 4, Panel A).
C. Yang et al. / Journal of Colloid and Interface Science 450 (2015) 388–395
A recent study showed that aggregations of 6 nm AuNPs on bacterial surface through hydrophobic interactions between proteins and AuNPs. And aggregation of 2 nm AuNPs on E. coli outer membrane were time-dependent and led to nanoparticle towers by resembling arrested precursors to the outer membrane vesicles [8], suggesting direct interactions between AuNPs and the cell surface. In the present study, no significant fluorescence change was observed after the surface proteins were removed (Fig. 4, Panels C and D), implying that proteins on cell surface did not involve in QDs/bacteria interactions. Actually, it was phospholipids molecules such as LPS on the membrane were demonstrated to involve in the adsorption and aggregation of QDs on E. coli cells. Since QDs used in the present study were modified with hydrophilic surface, it is not likely that hydrophobic interaction led to QDs/bacteria interactions. Therefore, it can be electrostatic attraction makes major contributions to QDs/bacteria interactions. The observation of time-dependent QDs aggregation on E. coli outer membrane (Fig. 3, Panel B, and Fig. 5) suggested that QDs may interact with E. coli cell surface in a manner similar to that of 2 nm AuNPs. In addition to LPS, positively charged QDs may also interact with other negatively charged molecules on bacterial surface, such as negatively charged phospholipid groups on bacterial inner membranes or eukaryotic cell membranes. Aggregation and fluorescence quenching of QDs may also occur upon interaction with these surface. Actually, significant fluorescence quenching did occur when QDs interacted with eukaryotic cells of Tetrahymena thermophile or ‘‘naked’’ cells of E. coli or B. subtilis whose cell wall were removed by lysozyme to expose their inner membrane (for details, see the Supporting Information). Therefore, positively charged QDs can be broadly used in interaction with both bacterial and eukaryotic cells. 5. Conclusions The present study reports the behavior and direct interactions between the cysteamine-stabilized CdTe QDs and bacterial cells. Distinct fluorescence quenching patterns were developed when Gram negative or Gram positive bacteria cationic interacted with QDs. Aggregation of QDs and fluorescence quenching was observed and related to chemical compositions and structures of bacterial cell envelopes. Negatively charged groups of molecules such as LPS may also play an important role in the interaction between hydrophilic cationic nanoparticles and bacterial cells. The potential application was also demonstrated in rapid discrimination of Gram-negative and Gram-positive bacteria. The present study may not only provide insight into behaviors of QDs on bacterial cell surfaces but also open a new avenue for designing and applying QDs as biosensors in microbiology, medicine, and environmental science. Further fluorescence-based detections and measurements of intact bacterial cells can be achieved based on the direct interactions between cationic QDs and bacterial cell surface. Acknowledgments This work is financially supported by ‘‘the Fundamental Research Funds for the Central Universities’’ (WHUT, 2013-Ia037). B.-L. Su also acknowledges the Chinese Central Government for an ‘‘Expert of the State’’ position in the program of ‘‘Thousand
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