Nanothermometry Measure of Muscle Efficiency - ACS Publications

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Nanothermometry Measure of Muscle Efficiency Suvra S. Laha, Akshata R. Naik, Eric R. Kuhn, Maysen Alvarez, Alyson Sujkowski, Robert J. Wessells, and Bhanu P. Jena* Department of Physiology, Wayne State University School of Medicine, Detroit, Michigan 48201, United States S Supporting Information *

ABSTRACT: Despite recent advances in thermometry, determination of temperature at the nanometer scale in single molecules to live cells remains a challenge that holds great promise in disease detection among others. In the present study, we use a new approach to nanometer scale thermometry with a spatial and thermal resolution of 80 nm and 1 mK respectively, by directly associating 2 nm cadmium telluride quantum dots (CdTe QDs) to the subject under study. The 2 nm CdTe QDs physically adhered to bovine cardiac and rabbit skeletal muscle myosin, enabling the determination of heat released when ATP is hydrolyzed by both myosin motors. Greater heat loss reflects less work performed by the motor, hence decreased efficiency. Surprisingly, we found rabbit skeletal myosin to be more efficient than bovine cardiac. We have further extended this approach to demonstrate the gain in efficiency of Drosophila melanogaster skeletal muscle overexpressing the PGC-1α homologue spargel, a known mediator of improved exercise performance in humans. Our results establish a novel approach to determine muscle efficiency with promise for early diagnosis and treatment of various metabolic disorders including cancer. KEYWORDS: Nanometer scale thermometry, muscle efficiency, disease detection, quantum dot

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dvances in thermometry1−8 have enabled precise determination of temperature although major challenges still remain at the nanometer scale in live cells. Possibilities range from understanding subcellular metabolism at nanometer resolution to the early detection of cancers,9 cardiac,10,11 and skeletal12 muscle disorders. In the past decade, zero-dimensional, semiconducting, nanometer crystals or quantum dots (QDs) have attracted considerable attention for their promise in physical, chemical, and biomedical applications, ranging from optoelectronics to medical diagnostics. 13−15 QDs with enhanced photostability and brightness are considered superior to standard organic dyes, green fluorescent proteins,7 fluorescent polymers6 and lanthanide-based chelates for the investigation of essential life processes at the nanometer scale.13−15 Recent studies report the potential use of QDs as nano- thermometers for probing local temperatures in biological systems.4,16,17 The emitted fluorescence intensity and the spectral wavelength shifts of the QDs are the two vital parameters for temperature sensing.17,18 Fluorescent intensity of QDs decreases following an increase in temperature. A red spectral shift is occasionally associated with a rise in temperature as well, altering the QD emission color.17 In most semiconductors, these temperature-dependent wavelength shifts have been attributed predominantly to electron− phonon interactions and to thermal expansion of the lattice.19 On the basis of these properties of QDs, studies2 report cadmium telluride (CdTe) QDs as excellent temperaturesensing nanoprobes with a precision of approximately 0.2 °C. Similarly, cadmium selenide (CdSe) QDs have been used as nanothermometers with a resolution of ∼1 °C depending on wavelength shift in spectroscopic studies.1 Although promising © XXXX American Chemical Society

thermometric approaches on live cells using QDs have been used in previous studies,4,19 they have been functionalized with tethers measuring several nanometers, precluding direct thermometric measurement of the target. The physical contact of QDs with a biomolecule, cellular compartment, cell, or tissue under study is therefore required for precise and meaningful thermometry. In this study, we conjugate CdTe QDs directly to our target muscles or muscle proteins to obtain a specific thermometric measurement, allowing us to precisely quantify muscle efficiency. Results and Discussion. Myosin, a molecular motor that binds and hydrolyses ATP required for muscle contraction and movement, is schematically represented in Figure 1A. Heat loss by a molecular motor while doing work provides a way of quantifying its efficiency. Changes in the performance of cardiac or skeletal muscles are an early indication of the health status of the tissue, providing an opportunity for timely diagnosis and intervention. Therefore, we have used monodispersed, core-type CdTe QDs averaging 2 nm in diameter (Figure 1B) as a novel tool capable of sensing a broad temperature range (24−42 °C; Figure 1C). Furthermore, these QDs are optimized to directly adhere to myosin molecules (Figure 1D,E) and quantitatively estimate the heat release from ATP hydrolysis (Figure 2). There is a temperature-dependent change in QD fluorescence, demonstrating decreasing intensity with increasing temperature with a thermal resolution of Received: December 7, 2016 Revised: January 17, 2017 Published: January 23, 2017 A

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RS myosin II and bovine cardiac (BC) myosin hydrolyze ATP (Figure 2). ATP hydrolysis was estimated and confirmed from the inorganic phosphate released (Supporting Figure S2). Equal amounts (0.22 μM) of RS and BC myosin proteins generated similar amounts of inorganic phosphate when exposed for 10 min to 3 mM ATP (absorbance 650 nm: 0.368 ± 0.052 (RS) and 0.366 ± 0.075 (BC)) (Figure 2G), demonstrating equal moles of ATP hydrolyzed by both motors. Interestingly, ATP hydrolysis by the BC myosin released a greater amount of heat and consequently a larger drop in fluorescence compared to RS myosin-associated CdTe QDs (Figure 2). Therefore, BC myosin is less efficient than the RS myosin as a molecular motor (Figure 2 and Supporting Information). To our knowledge, this is the first demonstration of the use of QDs in direct physical association with a ATPase molecular motor to determine thermal shifts when catalyzing ATP hydrolysis. Fluorescence microscopy on CdTe QD-associated RS and BC myosin proteins exposed to 3 mM ATP under similar experimental conditions, demonstrated a gradual time-dependent drop in fluorescent intensity for the RS myosin compared to BC (Figure 2A−C). Interestingly, within 15 s following ATP additio, BC myosin showed a relatively sharp decay in fluorescence (Figure 2B,C). In the presence of the nonhydrolyzable ATP analogue, adenylyl imidodiphosphate (AMPPNP), and myosin-(2′,7′-bis(2-carboxyethyl)-5-(and-6)-carboxyfluorescein acetoxymethyl ester) (BCECF is a pH sensitive dye in the presence of ATP), Myosin-QDs showed little timedependent loss of fluorescence, demonstrating minimal photobleaching during imaging, or possible pH changes upon ATP addition. Hence, the observed fluorescence drop when myosinQDs are exposed to ATP is primarily a thermal phenomenon. The calculated data presented in Figure 2C reveal that during the initial 15 s following ATP addition, the rate of fluorescence loss in the BC-QD sample is more than twice the rate of RSQD (rBC = −0.047 s−1, rRS = −0.021 s−1) (Figure 2B,C). The fluorescence decay for all three controls show similar behavior, as the slopes (r) are an order of magnitude lower than those corresponding to hydrolysis by RS and BC myosin (experimental). The intensity profiles when myosin proteins hydrolyze ATP shown in Figure 2C can also be fitted to an exponential decay relation, I ∼ e−λt, with λ being the decay constant, resulting in λRS and λBC values of 0.04 and 0.09 s−1, respectively (λBC ∼ 2λRS), reflecting greater heat loss by the BC myosin when hydrolyzing ATP. To further evaluate the efficiency of RS and BC myosin proteins during ATP hydrolysis, real-time continuous spectrofluorimetric measurements were performed (Figure 2D.E). Although the amount of hydrolyzed ATP is the same for both the myosin proteins under similar experimental conditions (Figure 2G), the potency and efficacy of loss in QD fluorescent intensity in the BC myosin is sharper and greater compared to RS (Figure 2E). Quantitative estimation in the initial 60 s following ATP addition to BC and RS myosin demonstrated a 20% greater heat loss by BC over RS (Figure 2F), signifying BC to be less efficient then RS. As observed in our fluorescent microscopy studies (Figure 2B,C), the relative rate of loss in fluorescence using spectrofluorimetry is similar in both controls (QD-RS myosin and QD-RS myosin-AMP-PNP), (Figure 2D,E; r = −0.002 s−1). Likewise, in the initial 15−20 s following ATP exposure, the fluorescent decay rate in the BC experimental (QD-BC myosin-ATP) is much sharper, nearly twice that of RS (rRS = −0.014 s−1, rBC = −0.027 s−1) (Figure 2E). The intensity profiles for both the RS and BC myosin

Figure 1. Thermometry on the molecular motor myosin using nanometer-scale sensors. (A) Schematic depicts bright green fluorescence of ∼2 nm sized cadmium telluride quantum dots (CdTe QDs) bound to myosin and the resulting decrease in QD fluorescence as a consequence of heat released from myosin-mediated ATP hydrolysis. (B) Size measurement of CdTe QDs using photon correlation spectroscopy (PCS) in a zeta sizer demonstrates an average particle size of ∼2 nm. (C) Relative fluorescent intensity of CdTe QDs dispersed in phosphate buffered saline (pH 7.4) at different temperatures. Fluorimetry shows the temperature-dependent decrease in fluorescent intensity of the QDs for four representative experiments (n = 4), demonstrating a temperature sensitivity of ∼1 mK (Figure S1). (D) PCS of purified rabbit skeletal muscle myosin demonstrates an average particle size of 24.36 nm. (E) PCS on QD-functionalized rabbit skeletal muscle myosin measures 28.21 nm with no free QDs in the suspension, hence demonstrating complete binding, critical in eliminating background fluorescence.

approximately 1 mK (Figure 1C and Supporting Figure S1). Standards for the estimation of inorganic phosphate released (Supporting Figure S2) and the determination of optimal ATP concentration used to measure ATP hydrolysis are available in the Supporting Information (Supporting Figure S3). Fluorescence of rabbit skeletal (RS) muscle myosin II-bound CdTe QDs drops monotonically when exposed to ATP (1−5 mM), showing no change in intensity beyond the 3 mM ATP concentration (Supporting Figure S3). Consequently, all of our experiments were performed using either 3 or 5 mM ATP. Using fluorescent microscopy and real-time spectrofluorimetric measurements, we have determined the drop in QD fluorescence as a consequence of heat released when purified B

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Figure 2. RS myosin is more efficient than BC myosin. (A) Fluorescent images of CdTe QDs associated with RS and BC myosin proteins at different time points following addition of 3 mM ATP. Calculations reveal an average of 5 QDs directly bound to a single myosin molecule (see Supporting Information). Note the rapid loss of fluorescence in the BC-associated QDs resulting from greater heat release following ATP hydrolysis. (B) Time-dependent drop in fluorescent intensity estimated from fluorescent micrographs in one of three representative experiments. (1) RS myosin-BCECF + 3 mM ATP (purple inverted triangles); (2) RS myosin-QDs (green squares); (3) BC myosin-QDs + 3 mM AMP-PNP (brown stars); (4) RS myosin-QDs + 3 mM ATP (red circles); and (5) BC myosin-QDs + 3 mM ATP (blue triangles). Arrowhead indicates ATP addition. (C) The slopes (r) indicate a time-dependent loss in fluorescent intensity following ATP addition. r1 and r2 indicate the rate of fluorescence decay in RS and BC myosin proteins during the initial 15−20 s and at later stages, respectively. In the initial 15−20 s, r1 for BC is more than doubled than in RS. Exponential fit (black lines) resemble first-order reaction kinetics of RS myosin-QDs and BC myosin-QDs in the presence of 3 mM ATP demonstrates that the exponential decay constant or first-order rate constant λ in BC is roughly double of RS. (D) Spectrophotometry demonstrates a time-dependent drop in fluorescence of CdTe QDs following ATP hydrolysis by myosin, similar to the observations made using fluorescent microscopy. Arrowhead indicates the addition of myosin-QDs. (1) RS myosin-BCECF + 3 mM ATP (purple inverted triangles); (2) RS myosinQDs (green squares); (3) BC myosin-QDs + 3 mM AMP-PNP (brown stars); (4) RS myosin-QDs + 3 mM ATP (red circles); and (5) BC myosinQDs + 3 mM ATP (blue triangles). (E) The rates of fluorescence loss in RS and BC myosin proteins during the initial 15−20 s and at later time points are similar to those observed using fluorescence microscopy. r1 for BC is twice as much for RS. Similarly, exponential fit (black lines) of RS myosin-QDs and BC myosin-QDs in 3 mM ATP, shows that the value of λ in BC is approximately twice as that for RS myosin. (F) Percent drop in fluorescence, as a function of time, when RS and BC myosin proteins are exposed to 3 mM ATP. Note the significant (p < 0.01) drop in timedependent fluorescent intensity of CdTe QDs associated with BC myosin over RS myosin. Data is presented as mean ± S (n = 3). (G) The inorganic phosphate (iP) release assays show that similar quantities of ATP (3 mM) is hydrolyzed by equal amounts (0.22 μM) of both RS and BC myosin proteins. (H) RS-associated QDs showing a spatial resolution of approximately 80 nm (red square).

recorded (Supporting Information). However, because a heat loss equivalent to 35 and 45 °C by the RS and BC myosin, respectively, is observed, this translates to an efficiency of 30% and 10% respectively for the two myosin types. In vertebrates, including humans, the transcriptional cofactor, peroxisome proliferator-activated receptor gamma coactivator 1-alpha (PGC-1α), is upregulated in response to endurance exercise training,20 and muscle-specific overexpression of PGC1α significantly improves exercise and aerobic capacity.21,22 A homologue of PGC-1α called spargel is present in Drosophila melanogaster whose function is similarly conserved.23,24 Musclespecific expression of spargel mimics the effects of exercise on various aspects of Drosophila physiology.25 To test the hypothesis that overexpression of spargel would potentiate efficiency of the fly muscle in the use of ATP, CdTe QDs were directly associated with isolated skeletal muscles obtained from wild type Drosophila melanogaster outcrossed genetic background control flies (y1w1;UAS-srl) and flies with musclespecific spargel overexpression (mef2> UAS-srl). Upon

proteins follow a typical exponential behavior (first-order reaction kinetics, λRS = 0.03 s−1 and λBC = 0.06 s−1), with greater amount of heat loss from the BC myosin-mediated ATP hydrolysis (Figure 2E). Therefore, these fluorimetric measurements are in good agreement with those obtained using fluorescent microscopy (Figure 2A−C). Using fluorimetry, we observed that the loss in fluorescent intensity of myosin-QDs in the first 60 s following ATP exposure of RS and BC (Figure 2F) is approximately 60% and 80%, respectively. This timedependent fluorescent intensity drop in CdTe QDs observed in Figure 1C translates to a temperature rise of approximately 35 and 45 °C in the RS and BC myosin, respectively, with a thermal sensitivity of ∼0.001 °C or 1 mK (Supporting Figure S1), and a spatial resolution of 80 nm (Figure 2H). Calculations (Supporting Information) of the complete hydrolysis of 2 nmoles of ATP correspond to a release of [(7.3 kcal/mol × 2 nmoles) = 14.6 μcal = (14.6 × 4.2) = 60 μJ] 60 μJ of energy (1 cal = 4.2 J). If the entire energy had been lost as heat, a 50 °C rise in temperature would have been C

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Figure 3. Expression of spargel in Drosophila melanogaster, a known homologue of the endurance exercise-induced vertebrate gene PGC-1α, results in improved efficiency of skeletal muscles in the fly. (A) The phase (left) and fluorescent image (center) of the skeletal muscles of Drosophila melanogaster at a resolution of 80 nm (red square, far right image). (B) The phase and fluorescent images of CdTe QD-associated skeletal muscles of Drosophila melanogaster, y1w1; UAS-srl (control) and mef2> UAS-srl (experimental) following exposure to 5 mM ATP at room temperature. Note the increased loss of fluorescence in the control tissue compared to the spargel-expressing experimental tissue. (C) Fluorescent intensity plot demonstrates significantly greater time-dependent drop in fluorescence of CdTe QDs associated with the control muscle (green triangles) over experimental muscle (red circles). Stars indicate statistical significance (n = 8). (D) Extent of ATP hydrolysis in control and experimental muscles, expressed as inorganic phosphate release. Absorbance recorded at 650 nm at 10 min following addition of 5 mM ATP indicates that the amount of iP released (nanomoles) is similar in both control and experimental muscles (n = 8).

s−1, rexp. = −0.004 s−1), almost twice as fast as in the spargelexpressing experimental, especially during the initial 30 s of ATP addition (Supporting Figure S4). Approximately equal amounts of inorganic phosphate released by both the control and experimental muscles following exposure to ATP (Figure 3D), signifies similar quantities of ATP hydrolysis by both muscle tissues. Furthermore, studies using microscopy demonstrate that exposure of isolated fly muscle to the myosin

exposure to 5 mM ATP, time-dependent fluorescent intensity changes were recorded using fluorescent microscopy (Figure 3A−C). Similar to purified myosin (Figure 2), the fluorescent image heat maps (Figure 3B) of control (y1w1; UAS-srl) and experimental (mef2> UAS-srl) fly skeletal muscle preparations following ATP hydrolysis demonstrate a greater heat loss in control muscles over experimental. The drop in CdTe QD fluorescence is relatively sharp in the control (rcont. = −0.007 D

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acquired from the diffusion coefficient.31,32 The intensity size distribution, which is obtained as a plot of the relative intensity of light scattered by particles in various size classes, was then calculated from a correlation function using built-in software. Preparation of Myosin and Myosin-QD Suspension. One milligrams of myosin (MYO) protein (Cytoskeleton, Inc., Denver, CO) was diluted in 100 μL of myosin resuspension buffer (15 mM Tris HCl of pH 7.5, 0.2 M KCl and 1 mM MgCl2) to obtain a final concentration of 10 mg/mL. Ten microliters of myosin protein (10 mg/mL) was incubated with 3 μL of CdTe QDs (5 mg/mL) for an hour to obtain MYOQDs, and the 100% binding of QDs to MYO was determined using the Zetasizer (Figure 1B,D,E). Graphs were plotted and analyzed using Origin 8.5. Measurement of Temperature from Fluorescent Intensity of CdTe QDs. The fluorescent intensity of the CdTe QDs as a function of temperature was estimated using a Hitachi F-2000 Fluorescence spectrophotometer (Figure 1C; Supporting Figure S1). Ten to fifteen microliters of CdTe QDs at a concentration of 5 mg/mL was resuspended in phosphate buffered saline (PBS) pH 7.4, and the temperature of the suspension was raised. Real-time measurement of temperature and fluorescent intensities of the QD suspension was estimated using a pinhead thermometer and a Hitachi U-2000 Spectrophotometer. Accordingly, fluorescent intensities at different temperatures were measured. The excitation wavelength was kept fixed at 350 nm (λex = 350 nm) for these  COOH functionalized CdTe core-type QDs (Sigma-Aldrich), and emission collected at ∼520−530 nm. Fluorescence Microscopy. Imaging was performed using the Axiovert 200 (Carl Zeiss, Munich, Germany), 10× (numerical aperture 0.5) and 100× (numerical aperture 1.4). One microliter of MYO-QDs (10:3) was applied to the surface of a glass slide, followed by an hour of incubation, and then washed (2×) in 10 vol of the myosin resuspension buffer. The sample was imaged prior to and following addition of 3 mM ATP. Images were captured at an interval of 30 s for a period of 3 min. Phosphate Standard and ATP Hydrolysis. The amount of free phosphate following ATP hydrolysis was estimated using a phosphate standard curve determined using published procedure.28 The standard curve was established using the CytoPhos reagent kit (Cytoskeleton Inc., Denver, CO). A 0.22 μM sample of both rabbit skeletal and bovine cardiac myosin molecules was exposed to 3 mM ATP (Supporting Figure S2). Incubation at 37 °C was maintained for 10 min. After 10 min, CytoPhos TM was added to terminate the reaction. Following an incubation of another 10 min at room temperature, the absorbance at 650 nm was recorded. Real-Time Fluorescent Spectroscopy. Real-time fluorescent spectroscopy was performed on free QDs and MYO-QDs in the presence and absence of ATP using a quartz cuvette in a Hitachi F-2000 Spectrophotometer (Hitachi, Tokyo, Japan). Time-dependent percent drop in fluorescence was then calculated from the data obtained online (Supporting Figure S3). Isolation of Drosophila melanogaster Skeletal Muscles. Skeletal muscles were isolated from Drosophila melanogaster, y1w1; UAS-srl (control) and mef2> UAS-srl (experimental). Briefly, male flies were immobilized dorsal side up with pins in a drop of phosphate buffered saline pH 7.4, and thoraces were opened with small scissors to allow separation of indirect flight muscle (IFM) with sharp tweezers. Dissected muscle was

II ATPase in hibit or 4-methy l- N-(p hen ylm eth yl)benzenesulfonamide (BTS),27−29 and to the vH-ATPase inhibitor Bafilomycin,30 demonstrate the near abolition of CdTe QD fluorescence drop in the presence of BTS as opposed to a fluorescent loss in the presence or absence of Bafilomycin (Supporting Figure S5). Similarly, spectrofluorimetric analysis of bovine cardiac myosin exposed to BTS demonstrates the near abolition of CdTe QD fluorescence drop as opposed to the control (vehicle) (Supporting Figure S6). These results (Supporting Figure S5 and S6) support that the heat release following exposure of fly muscle to ATP is primarily a result of the ATP hydrolysis by muscle myosin. Therefore, as hypothesized, the spargel-expressing fly muscle is found to be more efficient than wild type. Conclusion. These observations promise a platform for the early detection of various metabolic disorders such as muscular dystrophy, cardiovascular diseases, and cancer (Supporting Figure S7). Measurement of thermogenesis at the nanometer scale will allow detection of diseased cells at the single-cell level. As a result of increased glycolytic metabolism and higher metabolic rate, cancer cells exhibit higher temperatures,26 hence their detection using CdTe QDs. As a proof of concept, using nonmalignant MCF10A breast cancer cells, premalignant MCF10AneoT and TGF3B breast cancer cells, and malignant MCF10CA1h breast cancer cells, CdTe QD thermometry demonstrates a drop in fluorescent intensity in the MCF10AneoT and TGF3B cells and a much greater drop in the malignant MCF10CA1h cells compared to the nonmalignant MCF10A cells (Supporting Figure S7A). Similarly, CdTe QD thermometry using human head-neck cancer cells WSU12 (glycolytic) and UP154 (oxidative) demonstrated significant heat generation in the WSU12 cells compared to UP154, reflected in the loss of fluorescent intensity in WSU12 cells (Supporting Figure S7B). However, in order to further generalize the results on the use of CdTe QD in cancer detection, additional experiments on different forms of cancers are required and are in progress in the laboratory. The nature of CdTe QDs tightly adhered to a biological substrate such as a single biomolecule, cell, or a tissue has enabled precise thermometry at an unprecedented thermal resolution of 1 mK and a spatial resolution of 80 nm. Further refinement of the present ability to directly measure heat release from a chemical reaction will provide an improved comprehension of the dynamics of a wide range of molecular structure−function and disease detection in live cells. Materials and Methods. Suspension of CdTe QDs and Their Size Measurements. Ten milligrams of COOH functionalized CdTe core-type QDs with reported average size of 2.04 nm and λem = 520 nm (Sigma-Aldrich) were diluted in 2 mL of deionized double distilled and filtered water to obtain a 5 mg/mL stock solution and stored in the dark at 4 °C. Photon correlation spectroscopy (PCS) was used to study the size distribution by employing various dilutions of the suspension (Figure 1B). PCS measurements were performed in a Zetasizer Nano ZS, (Malvern Instruments, U.K.). The size distribution of QDs was determined using built-in software provided by Malvern Instruments. Prior to determination of the QD hydrodynamic radius, calibration of the instrument was performed using latex spheres of known size. In PCS, subtle fluctuations in the sample scattering intensity are correlated across microsecond time scales. The correlation function was calculated from which the diffusion coefficient was determined. Using Stokes−Einstein equation, hydrodynamic radius can be E

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Nano Letters removed directly onto the glass bottom coverslip of a Petri dish for imaging. Imaging Drosophila melanogaster Skeletal Muscles. Two microliters of CdTe QDs suspended in approximately 20−25 μL of the resuspension buffer were added to the Drosophila melanogaster skeletal muscle firmly fixed to the bottom of the Petri dish. After an hour of incubation, it was washed three times with the buffer to remove any unbound QDs. Subsequently, 5 mM ATP dissolved in 15 mM Tris-HCl pH 7.4 was carefully pipetted directly on the muscle at the bottom of the Petri dish. The muscle was then exposed to a blue monochromatic light source, following which both the fluorescence as well as the phase images were captured at regular intervals (30 s) for a period of 4 min. Estimating the Number of CdTe QDs Adhering to a Single Myosin Molecule. Calculations reveal that approximately 5 QDs are directly bound to a single myosin molecule. In our experiments, we incubated 10 μL of myosin protein (10 mg/ mL) with 3 μL of CdTe QDs (5 mg/mL). The molarity corresponding to 10 mg/mL of myosin is around 20 μM. Using the density of CdTe = 6.2 g/cm3, mass of CdTe QDs is 10 mg, the approximate size of a CdTe QD is 2 nm (Figure 1), and considering each QD to be spherical having a radius of 1 nm, the number of QDs in 10 mg = 3.85 × 1017. Therefore, the number of particles in 5 mg of QD sample should be 2 × 1017. So 3 μL of CdTe having a concentration of 5 mg/mL contains approximately 6 × 1014 QDs. Rabbit skeletal myosin II with a molecular weight of 520 kDa is comprised of 6.023 × 1023 molecules (Avogadro number), hence 10 mg of myosin has 1.2 × 1016 molecules. Therefore, 10 μL of myosin having a concentration of 10 mg/mL contains 1.2 × 1014 molecules, hence each myosin molecule is bound to an average of 5 CdTe QDs. Estimation of the Temperature Increase during ATP Hydrolysis. ATP hydrolyzes to ADP and Pi with the release of 7.3 kcal of energy per mole of ATP. From the plate reader assay, we have estimated almost equimolar amounts of ATP being hydrolyzed by both RS and BC myosin proteins. Relating these data to the phosphate standard curve, we found that roughly 2 nmoles of ATP has been hydrolyzed releasing (7.3 kcal/mol ×2 nmoles) = 14.6 μcal = (14.6 × 4.2) = 60 μJ of energy (1 cal = 4.2 J). In the fluorimeter experiment, 5 μL of MYO-QDs mixed in a ratio of 10:3 by volume was added to the cuvette (final volume = 350 μL) for performing the measurements. The MYO-QDs combination approximately contains 1 μL of CdTe QDs, which contributes to temperature sensing. Considering the density of CdTe = 5.86 g/cm3, the mass of the QDs in a volume of 1 μL = 5.86 × 10−3 g. Using the equation Q = cmΔT, where Q is the heat released per adsorbed QD, c is the specific heat, m is the mass of the material (CdTe in our case) and ΔT corresponds to the rise/ fall in temperature, we get ΔT = (60 × 10−6 J)/{(0.209) J/kg. °C × (5.86 × 10−3 × 10−3) kg} = 50 °C.

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ent decrease in fluorescence intensity of CdTe QD-RS myosin complex. Figure S4: Rate of loss in fluorescence intensity of CdTe QDs associated with Drosophila melanogaster skeletal muscles preparations. Figure S5: Microscopic assessment of relative fluorescence intensity of CdTe QDs associated with Drosophila melanogaster skeletal muscle upon ATP exposure. Figure S6: Relative fluorescence intensity of CdTe QDs associated with BC myosin in real-time, using continuous spectrofluorimetry. Figure S7: CdTe QDs thermometry performed on live nontransformed, pretransformed, and transformed (cancerous) cells, and in human head-neck cancer cell lines WSU12 (primarily use glycolytic metabolism) and UP154 (primarily use oxidative metabolism) (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Bhanu P. Jena: 0000-0002-6030-8766 Author Contributions

B.P.J. developed the idea, S.S.L., A.R.N., and B.P.J. designed experiments for the study. S.S.L. and A.R.N. conducted the experiments and analyzed the data. A.R.N. and E.R.K. conducted the experiments pertaining to the myosin-specific inhibitor BTS and analyzed and presented the data in Supporting Information. S.S.L. and B.P.J. wrote the manuscript. M.A., A.S., and R.J.W. generated the control (UAS-srl) and experimental (mef2> UAS-srl) Drosophila melanogaster cohorts and isolated skeletal muscle preparations for the study. All authors participated in discussions and proofread the manuscript. S.S.L and A.R.N contributed equally to this work. Notes

The authors declare the following competing financial interest(s): BPJ has filed for patent protection on the direct nano thermometry approach and technology and its use in disease detection.

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ACKNOWLEDGMENTS We thank Professor Hyeong-Reh Kim, Wayne State University, for providing nonmalignant (MCF10A), premalignant (MCF10AneoT and TGF3B), and malignant (MCF10CA1h) breast cancer cells and the human head-neck cancer cell lines WSU12 and UP154. This work was supported in part by the National Science Foundation Grant CBET1066661 (B.P.J), the WSU VP for Research Post-Doctoral Fellowship (S.S.L), and the WSU Interdisciplinary Biomedical Systems Fellowship (A.R.N).

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REFERENCES

(1) Li, S.; Zhang, K.; Yang, J.-M.; Lin, L.; Yang, H. Nano Lett. 2007, 7 (10), 3102−3105. (2) Maestro, L. M.; Jacinto, C.; Silva, U. R.; Vetrone, F.; Capobianco, J. A.; Jaque, D.; Solé, J. G. Small 2011, 7 (13), 1774−1778. (3) Kim, S. H.; Noh, J.; Jeon, M. K.; Kim, K. W.; Lee, L. P.; Woo, S. I. J. Micromech. Microeng. 2006, 16 (3), 526. (4) Yang, J.-M.; Yang, H.; Lin, L. ACS Nano 2011, 5 (6), 5067−5071. (5) Vetrone, F.; Naccache, R.; Zamarron, A.; Juarranz de la Fuente, A.; Sanz-Rodríguez, F.; Martinez Maestro, L.; Martín Rodriguez, E.; Jaque, D.; García Solé, J.; Capobianco, J. A. ACS Nano 2010, 4 (6), 3254−3258. (6) Okabe, K.; Inada, N.; Gota, C.; Harada, Y.; Funatsu, T.; Uchiyama, S. Nat. Commun. 2012, 3, 705.

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S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.nanolett.6b05092. Figure S1: Estimation of thermal resolution from fluorescent intensity of CdTe QDs. Figure S2: Inorganic phosphate standard curve. Figure S3: ATP dose-dependF

DOI: 10.1021/acs.nanolett.6b05092 Nano Lett. XXXX, XXX, XXX−XXX

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DOI: 10.1021/acs.nanolett.6b05092 Nano Lett. XXXX, XXX, XXX−XXX