Biophysical Journal Volume 100 February 2011 L19–L21 L19

Microsecond Single-Molecule Tracking (msSMT)

Stefan Semrau,* Anna Pezzarossa, and Thomas SchmidtPhysics of Life Processes, LION, Leiden University, The Netherlands

ABSTRACT Here we report on a method to track individual molecules on nanometer length and microsecond timescales usingan optical microscope. Our method is based on double-labeling of a molecule with two spectrally distinct fluorophores andilluminating it with laser pulses of different wavelengths that partially overlap temporally. We demonstrate our method by usingit to resolve the motion of short DNA oligomers in solution down to a timescale of 100 ms.

Received for publication 14 July 2010 and in final form 20 December 2010.

*Correspondence: semrau@mit.edu

Editor: David E. Wolf.

� 2011 by the Biophysical Society

doi: 10.1016/j.bpj.2010.12.3721

A persistent challenge in biophysics is measuring themotion of single biomolecules on short length scales andtimescales (1). Recent advances in superresolution tech-niques (2–5) have pushed the resolution limit of opticalmicroscopes down to the nanometer range. However, thesetechniques are severely limited in their temporal resolutionbecause they either require scanning the sample (as in stim-ulated emission depletion microscopy) or acquiring stochas-tically occurring signals (as in stochastic reconstructionmicroscopy) (STORM) and photoactivation localizationmicroscopy (2). Hence, they cannot access the fast dynamicinteraction of mobile biomolecules with nanometer-sized structures. For example, the interaction of a lipidmolecule diffusing in a cell membrane (diffusion coefficientD z 1 mm2/s) with a membrane substructure that has acharacteristic length L of ~50 nm occurs on a timescaletz L2/4Dz 1 ms. Consequently, observing the interactionof molecules with nanometer-sized structures requires tech-niques with submillisecond temporal resolution.

Here we present what is, to our knowledge, a new camera-based, thus highly parallel, single-molecule tracking (SMT)method that fulfills this requirement. The two major factorswhich determine the temporal resolution of camera-basedSMT (6) are the acquisition speed of the charge-coupleddevice (CCD) camera and the illumination time necessaryto detect the optical probe. During the illumination time,the probe must emit enough photons to be detected abovethe read-out noise of the camera. At the same time, theread-out noise generally increases with the read-out speedof the camera. Consequently, SMT on microsecond time-scales could so far only be realized with colloidal gold probes(7) or highly elaborate optical detection schemes (8–10).Although colloidal probes are bright and optically stable,they are not optimal for biological applications because oftheir large size and unspecific binding issues. Existing fastdetection schemes for single molecules can only track onemolecule at a time (8–10) or have a small field of view (10).

In contrast, our method uses small fluorophores in combi-nation with a simple detection scheme—a slow-scan, lowreadout-noise CCD camera system—to achieve micro-

second single-molecule tracking (msSMT). msSMT iscomparable to STORM in that it uses two fluorescent probeson a single molecule. In contrast to STORM, these probesare placed so far apart that no energy transfer can take placebetween them and thus the two are excited separately.

The emitted signal of the twofluorophores is collected ontoseparate regions of a CCD chip with the help of a dichroicwedge (see Fig. S1 in the Supporting Material). In this way,the two fluorophores are separately excited and their fluores-cent signals recorded independently of each other. Therefore,we can resolve the position of a single molecule with a nano-meter resolution at two different points in time.

To demonstrate our method, we used short DNA-oligonu-cleotides in solution that were labeled with two fluorophores,Cy3B and ATTO647N. These probes were then separatelyilluminated by two lasers at wavelengths of 514 nm and639 nm, respectively, during a certain illumination time till.The pulses of the two lasers were temporally offset by a smalltime lag Dt (see Fig. 1 a). In principle, we can make Dt arbi-trarily small. The actual temporal resolution achievable inpractice is primarily determined by the molecule’s diffusioncoefficient, the accuracy with which we can determine themolecular positions, and the total number of single-moleculepositions measured.

Because the illumination time was finite, we did notmeasure the actual positions rðtÞ and rðt þ DtÞ of the mole-cule. Instead, our method measured the positions

rðtÞ and rðt þ DtÞaveraged over the illumination time till. Due to this temporalaveraging, the mean squared-displacement (MSD) calculatedfrommeasuredpositionswas different from the actualMSDofthe molecule (11). For ergodic and stationary processes thereis, however, a simple relationbetween the observed (averaged)MSDðDtÞand the actual MSD of the molecule

FIGURE 1 Illumination scheme and position determination. (a)

Laser pulses of duration till are temporally offset by a time lagDt.

(b) Raw image from the Cy3B channel. (c) Raw image from the

ATTO647N channel. Dt ¼ 0 ms. Scale bar is 5 mm. (d) Correction

for chromatic aberration. The scheme shows positions in the

Cy3B (green dots) and ATTO647N (red dots) channel which

correspond to the same physical positions. (e) Identified

single-molecule signals from the images shown in panels

b and c, corrected for chromatic aberration.

L20 Biophysical Letters

MSDðDtÞ ¼Dðrðt þ DtÞ � rðtÞÞ2

E

¼ 1

t2ill

Z 0

�till

dt0Z �t0

�till�t0d3ðMSDðDtþ3Þ�MSDð3ÞÞ:

ð1ÞBecause the microscope projects the three-dimensional

movement of the molecules in solution onto a two-dimen-sional plane,

MSDðDtÞ ¼ 4DDt:

For regular diffusion, Eq. 1 was evaluated analytically (see

the Supporting Material for the derivation of the generalrelation and the analytical evaluation).

MSDðDtÞ ¼ 4D�

n�Dt2

till� 1

3

Dt3

t2ill

�for Dt%till

�Dt � 1

3till

�for DtRtill

:(2)

Below we will show that the measured MSDðDtÞ indeed

behaved as predicted by Eq. 2.

The calculation of amolecule’sMSD requires the determi-nation of its position. In our experiments, we chose a lowdensity of molecules such that molecules were resolved indi-vidually and their position determined with nm positional

Biophysical Journal 100(4) L19–L21

accuracy (6). Fig. 1 illustrates the process of position determi-nation. Fig. 1, b and c, shows typical raw signals from severalmolecules dual-labeled with Cy3B (Fig. 1 b) andATTO647N(Fig. 1 c). The positions of the fluorophores were determinedby fitting two-dimensional Gaussians approximating thepoint-spread function of themicroscope. The positional accu-racy achievedwas between 40 and 50 nm (see the SupportingMaterial). Because the signals from the two fluorophoreslabeling the singlemolecule had to be correlated, eliminatingchromatic aberration was important. We established thecorrespondence between the two signals by using fluorescentbeadswhichwere observable in both channels. Fig. 1 d showsthe corrective shifts which were applied to the signals in thegreen (or red) channel, respectively. Fig. 1 e shows thesingle-molecule positions determined from Fig. 1, b and c,and corrected for chromatic aberration. Note that due to thefinite labeling efficiency and bleaching, some moleculescarried no or only one of the labels.

To determine the MSD of the molecules, we correlatedthe positions that were measured in the two detection chan-nels by particle image cross-correlation spectroscopy(PICCS, see the Supporting Material), an extension ofparticle image correlation spectroscopy (PICS) (12).Because PICCS relies on highly accurate detection ofsingle-molecule positions, our method is not diffraction-limited and unaffected by autofluorescence and noise, bothof which are common problems faced in the conventionalimaging methods. Fig. 2 shows a comparison of MSDsobtained experimentally under different solvent and illumi-nation conditions. Equation 2 fits the experimental data inall cases. The inset table in Fig. 2 compares the diffusioncoefficients determined from the fit. As to be expected,the diffusion coefficient decreased with increasing amountof dextran T500 corresponding to an increased viscosity.

Compared to the situation without the dextran, the diffu-sion coefficient decreased by a factor of 3.4 5 0.9 in thesolution with 5% dextran T500 and by a factor of 5.8 51.0 in the solution with 10% dextran T500 (see inset tablein Fig. 2 b). We confirmed our results by independent fluo-rescence correlation spectroscopy measurements (see theSupporting Material) of the same DNA construct in thesame solvents in which we found factors of 3.20 5 0.04and 6.055 0.06 between the corresponding diffusion times,respectively. Interestingly, the diffusion coefficient did notscale with the bulk kinematic viscosity which increased bya factor of 6.2 5 0.1 in the solution with 5% dextranT500 and by a factor of 20.9 5 0.2 in the solution with10% dextran T500, as determined by bulk viscositymeasurements. This deviation probably reflects the factthat solvent and solute molecules were comparable in sizefor which the Stokes-Einstein relation is invalid.

For nonoverlapping laser pulses (Dt > till), the observedMSD was shifted down by (4/3) Dtill (see Fig. 2 a), in agree-ment with earlier results (11). For overlapping laser pulses(Dt % till), the MSD was clearly nonlinear and approached

FIGURE 2 Experimentally obtained MSDs of DNA with different

solvent and illumination conditions. In phosphate-buffered

saline (PBS), till ¼ 1 ms (black), in 5% dextran T500/PBS; till ¼3 ms (blue), in 5% dextran T500/PBS; till ¼ 5 ms (cyan), in 10%

dextran T500/PBS; till ¼ 3 ms (red), in 10% dextran T500/PBS;

and till ¼ 5 ms (orange). Equation 2 was fit to the data for each

experimental condition (solid lines). (Vertical dashed lines) Illu-

mination time for the respective experiment. (Error bars) Stan-

dard deviations of Monte Carlo simulations performed with the

measured parameters (see the Supporting Material). (a) Compar-

ison of all conditions, linear timescale. Above till, all MSDs are

linear. (b) Comparison of the measurements in dextran T500 at

short time lags Dt, logarithmic timescale. Below till, the MSDs

show the predicted asymptotic behavior. (Inset) Measured diffu-

sion coefficients for all conditions. The reported errors are the

errors of the fit to Eq. 2. (c) MSD on short time lags for the

measurement in PBS. The MSD has data points for Dt < till ¼ 1

ms, which are significantly different from 0 mm2.

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0 forDt/ 0. The temporal dependence followed the predic-tion given by Eq. 2 (see Fig. 2 b). Most importantly, as shownin Fig. 2 c, we were able to measure MSD curves down toDt ¼ 100 ms, significantly faster than reported so far, whichproved that we extended single-molecule tracking experi-ments to the microsecond regime.

Our method may be further improved by eliminating theneed for double labeling. Given that the fluorescent probe isbright enough, the emitted light could be split between twocameras that can be individually exposed. If the exposure ofthe two cameras is offset by a lag time Dt, the same kind ofanalysis as described here can be performed with onlya single fluorescent label.

Finally, msSMT can be easily adapted for measurementsin cells. For example, a membrane receptor could be labeledwith two genetically encoded tags, one at a cytosolicdomain, and one extracellular. The separation of the twolabels by the cell membrane would prevent energy transferbetween the fluorophores. Because only low laser intensitiesare required, the method is suitable for live cell recordings.Consequently, our method will likely aid in providing whatwe believe will be new insights into the dynamic behavior ofmolecules in cells on very short length- and timescales.

SUPPORTING MATERIAL

Supporting methods and theoretical derivations are available at http://www.

biophysj.org/biophysj/supplemental/S0006-3495(10)05294-X.

ACKNOWLEDGMENTS

The authors thank Ruth Buning for help with the fluorescence correlation

spectroscopy measurements, John van Noort for the DNA construct, Martin

van Hecke for help with the bulk viscosity measurements, and Hyun Youk

for critical reading of the manuscript.

This work is part of the research program of the Stichting voor Fundamen-

teel Onderzoek der Materie (FOM), which is financially supported by the

Nederlandse Organisatie voor Wetenschappelijk Onderzoek (NWO) within

the program on Material Properties of Biological Assemblies (under grant

No. FOM-L1707M).

REFERENCES and FOOTNOTES

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3. Rust, M. J., M. Bates, and X. Zhuang. 2006. Sub-diffraction-limitimaging by stochastic optical reconstruction microscopy (STORM).Nat. Methods. 3:793–795.

4. Betzig, E., G. H. Patterson, ., H. F. Hess. 2006. Imaging intracellularfluorescent proteins at nanometer resolution. Science. 313:1642–1645.

5. Hess, S. T., T. P. K. Girirajan, and M. D. Mason. 2006. Ultra-high reso-lution imaging by fluorescence photoactivation localization micros-copy. Biophys. J. 91:4258–4272.

6. Schmidt, T., G. J. Schutz, ., H. Schindler. 1996. Imaging of singlemolecule diffusion. Proc. Natl. Acad. Sci. USA. 93:2926–2929.

7. Tomishige, M., Y. Sako, and A. Kusumi. 1998. Regulation mechanismof the lateral diffusion of band 3 in erythrocyte membranes by themembrane skeleton. J. Cell Biol. 142:989–1000.

8. McHale, K., A. J. Berglund, and H. Mabuchi. 2007. Quantum dotphoton statistics measured by three-dimensional particle tracking.Nano Lett. 7:3535–3539.

9. Cang, H., C. S. Xu, and H. Yang. 2008. Progress in single-moleculetracking spectroscopy. Chem. Phys. Lett. 457:285–291.

10. Sahl, S. J., M. Leutenegger, ., C. Eggeling. 2010. Fast moleculartracking maps nanoscale dynamics of plasma membrane lipids. Proc.Natl. Acad. Sci. USA. 107:6829–6834.

11. Goulian, M., and S. M. Simon. 2000. Tracking single proteins withincells. Biophys. J. 79:2188–2198.

12. Semrau, S., and T. Schmidt. 2007. Particle image correlation spectros-copy (PICS): retrieving nanometer-scale correlations from high-density single-molecule position data. Biophys. J. 92:613–621.

Biophysical Journal 100(4) L19–L21