Name: luminescence resonance energy transfer to study conformational changes in membrane proteins expressed in mammalian cells
Uploaded: 2014-09-16T14:00:00.000+00:00
Duration: 8 min 31 s
Description: Watch this Scientific Journal Video about Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells at JoVE.com

This work was supported by National Institutes of Health Grant GM094246, the American Heart Association Grant 11GRNT7890004, and the National Science Foundation Grant MCB-1110501.

1. Create the Construct Containing the Protein of Interest
Clone the gene expressing the protein of interest into a suitable vector. Use vectors such as the pcDNA series or pRK5 as they are well suited for expression in mammalian systems like HEK293 and CHO cells.
2. Select the Sites on the Protein to be Tagged with the Fluorophores
<ol>
	<li>Choose labeling sites that are able to reflect possible conformational changes within the protein. If possible, use a crystal structure of the protein or of a homologous protein to help make this determination. If there is no crystal structure available, use online software to predict the possible structure of the protein and hence receive insight into appropriate sites.</li>
	<li>Choose residues such that the side chains of the selected residues are surface exposed and accessible to the fluorophores so that the protein can be labeled.
	<ol>
		<li>Choose residues that are not critical to protein function, for example sites that are not involved in ligand binding.</li>
		<li>In introducing cysteine mutation, give preference to sites that are similar, for example serine, that would cause minimal perturbation to the protein structure.</li>
		<li>After choosing the labeling sites, make the mutations to ensure that the protein only gives LRET from these intended sites. Use standard site-directed mutagenesis protocols to mutate away non-disulfide-bonded cysteines that could potentially bind to maleimide-conjugated fluorescent tags. Do not mutate away disulfide-bonded cysteines and buried, free cysteines since they should not react with fluorescent dyes in the folded form of the protein.</li>
	</ol>
	</li>
	<li>Introduce cysteine residues in the desired sites by making point mutations using standard site-directed mutagenesis protocols.</li>
	<li>Include a protease cleavage site that can specifically cleave off one of the cysteines from the protein. If the protein sequence allows for it, introduce the site by conservatively mutating the protein to have a thrombin (recognition sequence LVPRGS) or Factor Xa (recognition sequence IDGR or IEGR) sequence near the introduced cysteine and accessible to protease cleavage; otherwise the tetra- or hexa-peptide sequence may be inserted as a whole. Choose the site such that upon cleavage one of the introduced cysteines dissociates from the rest of the protein&#8212;in certain cases, this may require two cleavage sites flanking a mutated cysteine.</li>
</ol>
3. Test the Expression and Functionality of the Protein
<ol>
	<li>Perform a western blot to confirm expression of the mutated protein.</li>
	<li>Perform a functional assay of the protein to ensure that the mutations have altered the protein function only minimally, if at all, to avoid studying conformational changes of a dysfunctional protein. 
	NOTE: Since all proteins have different functions, there is no single functional assay that is used specifically for LRET; however, some examples of functional assays include enzyme activity assays for enzymes, ligand-binding assays for receptors, and electrophysiology studies for ion channels.</li>
</ol>
4. Select the Fluorophores to be Used
Select fluorophores based on the expected distance range being measured such that the range is between 0.5-1x&#160;the R0 of the fluorophore pair. 
NOTE: This allows for an easier subtraction of the background, which typically has much longer lifetimes. For example, if the expected distance range being measured is around 35 &#197;, an appropriate fluorophore pair to use would be terbium chelate as the donor and Alexa 594 as the acceptor, because the R0 for this pair is 53 &#197; (Table 1).
5. Express the Protein by Transiently Transfecting the Required Amount of Mammalian Cells
Transiently transfect the protein of interest into the chosen mammalian cells using any of the common transfection reagents. Typically, use four 10-cm dishes per LRET experiment for HEK and CHO cell lines; however, this amount may vary depending on protein expression, stability, etc. Allow the cells to express the protein for 36-48 hr before harvesting.
6. Label the Proteins
<ol>
	<li>Detach cells from the culture dish. Detach HEK cells by simply pipetting buffer against the bottom of the dish. Detach CHO cells with a cell scraper, washing off the media with extracellular buffer.</li>
	<li>Collect the cells by centrifugation at 1,100 x g for 3 min. Use these same centrifuge settings for collecting cells after labeling, as well as after the subsequent washes.</li>
	<li>Suspend cells in 3 ml of extracellular buffer, then add donor and acceptor fluorophores in equimolar amounts up to a final concentration of 100-300 nM. Incubate on a rotator for 1 hr at RT.</li>
	<li>Wash these labeled cells 3-4x&#160;with extracellular buffer to remove unbound fluorophores, then suspend these cells in extracellular buffer (usually 2 ml) for LRET measurements.</li>
	<li>As a control, label a separate set of cells with only donor fluorophore (terbium chelate) without addition of any acceptor fluorophore. 
	NOTE: Data from these donor-only experiments is necessary to complete analysis. These experiments can be done on the same or different days.</li>
	<li>Again, perform functional validation, this time with labeled mutant protein, as the labeling process may also interfere with function depending on the site used for labeling. 
	NOTE: These experiments can be done on the same or different days.</li>
</ol>
7. Set up the LRET Experiment
<ol>
	<li>Place the resuspended cells in a quartz cuvette with a minimum volume of 1 ml.</li>
	<li>Turn on the computer and the instrument and adjust the parameters of the data acquisition program accordingly.
	<ol>
		<li>Set the excitation wavelength to the absorbance range of the donor fluorophore (330-340 nm works well for terbium chelate).</li>
		<li>Set the emission wavelength appropriately, keeping in mind that the acceptor emission varies based on the acceptor used. Importantly, select a detection wavelength that measures only the acceptor emission and does not include any bleed-through from the donor&#8217;s emission. For example, use a wavelength of 565 nm for Alexa 555 as an acceptor Table 1. For donor-only measurements, in which protein was labeled only with donor but without acceptor, use a wavelength of 545 nm for measuring terbium chelate emission.</li>
		<li>Set the length of emission detection to be at least three times the expected LRET lifetime to ensure that a long-lifetime component will not be missed.</li>
	</ol>
	</li>
	<li>Perform the scan. Do at least three scans of 99 sweeps to ensure consistency of results. Save the results as a text file (*.txt).</li>
	<li>To measure conformational changes of a protein with regard to different conditions (such as the addition of a ligand), alter those conditions and again perform at least three scans of 99 sweeps each on the same sample under this new condition. If studying the effects of glutamate on the conformation of glutamate receptors, for example, add glutamate to 1 mM to the same sample scanned in step 7.3, and then scan again.</li>
	<li>Add up to five units of the appropriate protease and take scans continually until cleavage is complete and no further change is seen in the lifetime for three successive scans. Usually, cleavage is complete in two to 3 hr. If a Factor Xa cleavage site is being used for protease cleavage, for example, add 3 &#181;l of Factor Xa to the sample, and scan every 30 min for 3 hr.</li>
</ol>
8. Analyze the Data Obtained
<ol>
	<li>Open the data analysis software.</li>
	<li>Load the fluorescence lifetime data by using Import ASCII to open the text files. Load all the replicates as well as the final background measurements, into one file.</li>
	<li>Average the data from all the scans for each experimental condition to get the final data. To do so, highlight the columns that contain the individual trials, then under the Data menu in the menu bar, click on Statistics on Rows to display the average fluorescence intensities from the trials, as well as the standard deviation and standard error.</li>
	<li>Plot the average values as a line graph with fluorescence intensity on the Y-axis and lifetime in microseconds on the X-axis to create a curve that represents the lifetime of the sensitized emission of the acceptor. Change the Y-axis of the plot to a log scale, to allow for better visualization of the data.</li>
	<li>Repeat step 8.3 on the background measurement data, averaging the final scans that show overlap indicating complete cleavage.</li>
	<li>Display the mean background data on the same plot that contains the averaged raw data. To do so, open the Layer Control dialog box found under the Plot menu. Then, transfer the data containing the background mean into the list of Layer Contents.</li>
	<li>Align the mean background data to the mean raw data. To do so, use the Simple Math function under the Math menu to multiply or divide the background data as needed until the tail end of the background overlaps with the tail end of the raw data. 
	NOTE: At this tail end, the lifetime from the protein of interest should have already completely decayed away; what is aligned is simply the background signal present both before and after protease cleavage.</li>
	<li>Subtract the aligned background signal from the initial raw LRET signal, again using the Simple Math function.</li>
	<li>Fit the data to an exponential decay (single or multiple exponentials, depending on the sites and the experimental design).
	<ol>
		<li>Set the start boundary for fitting to begin after the end of the laser pulse. Use this same startpoint for all subsequent curve fittings.</li>
		<li>In the Select Fitting Function dialog box, select exponential decay so as to fit the lifetime to the following equation for a single exponential decay: 
		<img alt="Equation 4" fo:content-width="2in" src="/files/ftp_upload/51895/51895eq4.jpg" width="200" />&#160;Eq. 4 
		where, y represents the fluorescence intensity, y0 represents the background intensity due to noise from the system, A1 is the signal intensity, t is the fluorescence lifetime, x is the time, and x0 is the time offset.</li>
		<li>Fix x0 to 0, start the Fitting Function, and fit the data. 
		NOTE: If the experiment is designed to have signal only from one pair of sites, the resulting data should easily be fit by a single exponential decay8. The residual of the fit is a good method to determine the goodness of the fit and if additional decay lifetimes are required.</li>
	</ol>
	</li>
	<li>Use the lifetimes obtained from the data to calculate the distance between fluorophores using the F&#246;rster equation (a rearrangement of Equations 1 and 2 above): 
	<img alt="Equation 5" fo:content-width="2in" src="/files/ftp_upload/51895/51895eq5.jpg" width="200" />&#160;Eq. 5 
	NOTE: All the variables are as mentioned above with &#964;DA having been measured as the sensitized acceptor emission lifetime. Further details and examples on measurements of R0 and R can be found elsewhere7,9,10.</li>
	<li>Repeat the experiment and data analysis at least three times to ensure reproducibility. Inconsistency between individual repetitions of the same measurement calls into question the validity of the data; use additional control experiments or repetitions. Calculate errors in measurement using the free Gustavus Error analysis calculator (developed by Dr. Thomas Huber) or a similar system that propagates the error in the fits of the lifetime.</li>
</ol>

<ol>
	<li>Selvin, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+and+biophysical+applications+of+lanthanide-based+probes.">Principles and biophysical applications of lanthanide-based probes.</a> Annual Review of Biophysics and Biomolecular Structure. 31, 275-302 (2002).</li><li>Stryer, L., Haugland, R. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Energy+transfer:+a+spectroscopic+ruler.">Energy transfer: a spectroscopic ruler.</a> Proceedings of the National Academy of Sciences of the United States of America. 58, 719-726 (1967).</li><li>Stryer, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+energy+transfer+as+a+spectroscopic+ruler.">Fluorescence energy transfer as a spectroscopic ruler.</a> Annual Review of Biochemistry. 47, 819-846 (1978).</li><li>Selvin, P. R., Hearst, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Luminescence+energy+transfer+using+a+terbium+chelate:+improvements+on+fluorescence+energy+transfer.">Luminescence energy transfer using a terbium chelate: improvements on fluorescence energy transfer.</a> Proceedings of the National Academy of Sciences of the United States of America. 91, 10024-10028 (1994).</li><li>Chen, J., Selvin, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thiol-reactive+luminescent+chelates+of+terbium+and+europium.">Thiol-reactive luminescent chelates of terbium and europium.</a> Bioconjugate Chemistry. 10, 311-315 (1999).</li><li>Ge, P., Selvin, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thiol-reactive+luminescent+lanthanide+chelates:+part+2.">Thiol-reactive luminescent lanthanide chelates: part 2.</a> Bioconjugate Chemistry. 14, 870-876 (2003).</li><li>Gonzalez, J., Rambhadran, A., Du, M., Jayaraman, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=LRET+investigations+of+conformational+changes+in+the+ligand+binding+domain+of+a+functional+AMPA+receptor.">LRET investigations of conformational changes in the ligand binding domain of a functional AMPA receptor.</a> Biochemistry. 47, 10027-10032 (2008).</li><li>Ramaswamy, S. S., Maclean, D. M., Gorfe, A. A., Jayaraman, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Proton+mediated+conformational+changes+in+an+Acid+Sensing+Ion+Channel.">Proton mediated conformational changes in an Acid Sensing Ion Channel.</a> J. Bio. Chem. 288, (2013).</li><li>Rambhadran, A., Gonzalez, J., Jayaraman, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conformational+changes+at+the+agonist+binding+domain+of+the+N-methyl-D-aspartic+acid+receptor.">Conformational changes at the agonist binding domain of the N-methyl-D-aspartic acid receptor.</a> The Journal of Biological Chemistry. 286, 16953-16957 (2011).</li><li>Rambhadran, A., Gonzalez, J., Jayaraman, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subunit+arrangement+in+N-methyl-D-aspartate+(NMDA)+receptors.">Subunit arrangement in N-methyl-D-aspartate (NMDA) receptors.</a> The Journal of Biological Chemistry. 285, 15296-15301 (2010).</li><li>Kokko, T., Kokko, L., Soukka, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Terbium(III)+chelate+as+an+efficient+donor+for+multiple-wavelength+fluorescent+acceptors.">Terbium(III) chelate as an efficient donor for multiple-wavelength fluorescent acceptors.</a> Journal of Fluorescence. 19, 159-164 (2009).</li></ol>

The authors have nothing to disclose.

LRET is a powerful technique that allows scientists to measure distances between domains within a single protein as well as between subunits in a multimeric protein. As such, LRET is well-suited to examining the conformational changes and dynamics of proteins or other macromolecules. The above protocol should allow the properly equipped lab to easily test their hypotheses; however, there are many common sources of error that may plague the new investigator. If little or no LRET signal is seen, first check the wavelength ...

Luminescence Resonance Energy Transfer (LRET) is a derivative of the well-known Fluorescence Resonance Energy Transfer (FRET) technique1. Similar to FRET, LRET can be used to measure distances and distance changes between donor and acceptor fluorophores attached to specific sites on the protein of interest within the range of 10-100 &#197;1-3. The principles of LRET are also similar to FRET in that resonance energy transfer occurs between two proximal fluorophores when the emission spectrum of the donor fluorophore overlaps with the absorption spectrum of the acceptor fluorophore. The efficiency of this transfer is related to the distance between the two fluorophores by the following equation:
<img alt="Equation 1" fo:content-width="1.2in" src="/files/ftp_upload/51895/51895eq1.jpg" width="120" />&#160;Eq. 1
where R is the distance between the two fluorophores, E is the efficiency of energy transfer, and R0, discussed below, is the F&#246;rster radius for the fluorophore pair, i.e. the distance at which efficiency of transfer is half-maximal. From this equation, one can see that efficiency is related to the magnitude of the distance raised to the inverse sixth power1. It is this inverse sixth power dependence that allows for FRET and LRET measurements to be exquisitely sensitive even to small distance changes when near the R0 of the FRET pair. The ability to specifically label desired sites on proteins or other macromolecules allows one to take advantage of this sensitivity to monitor conformational changes.
When compared to FRET, which uses conventional organic dye molecules, LRET offers additional advantages. In LRET, instead of using an organic dye as the donor fluorophore, a lanthanide series cation, typically Tb3+ or Eu3+, is used1,4-6. Fluorophores that fall under this category, e.g., terbium chelate, are also very versatile in that they can be used with a wide range of acceptor fluorophores. This flexibility is made possible because the emission spectra of chelated lanthanides contain multiple sharp emission peaks, allowing for a single species of donor fluorophore to be used with one of a wide variety of acceptor fluorophores. Thus, sensitized acceptor emission can be detected without any fear of contaminating bleed-through from donor emission5. The experimenter selects the specific acceptor based on the expected distance between the two fluorophores (Figure 1 and Table 1). In these chelated lanthanide fluorophores, the metal ion is chelated by a molecule that contains an antenna group that sensitizes the normally poorly-absorbing lanthanide to excitation as well as a bioreactive functional group to tether the ion to a specific functional group on the macromolecule1,5,6. Once excited, lanthanides relax to the ground state via the release of photons with a decay rate in the millisecond range. Because the decay is neither a singlet-to-singlet relaxation nor a triplet-to-singlet relaxation, the emission of photons cannot properly be called fluorescence or phosphorescence, but is more properly termed luminescence1. The long decay of lanthanide luminescence greatly helps in lifetime measurements. Lifetime measurements can then be used to determine efficiency via the following relation:
<img alt="Equation 2" fo:content-width="1.2in" src="/files/ftp_upload/51895/51895eq2.jpg" width="120" />&#160;Eq. 2
where, E is the efficiency of transfer, &#964;D is the lifetime of the donor (chelated lanthanide) when not participating in energy transfer, and &#964;DA is the lifetime of the donor when participating in energy transfer with the acceptor. With LRET, &#964;DA can also be measured as the lifetime of the sensitized acceptor emission because terbium&#8217;s lifetime is so much greater than an organic acceptor fluorophore. The acceptor emits with the same lifetime as its inciting excitation (donor lanthanide), and any contribution to the lifetime from the acceptor&#8217;s own intrinsic fluorescence lifetime is relatively negligible. By measuring the sensitized emission rather than donor emission, we also eliminate the need to ensure labeling at exactly a 1:1 ratio of donor to acceptor. Protein can instead be labeled simultaneously with both acceptor and donor fluorophores. A heterogeneously labeled population will result, but double-donor labeled proteins will not emit in the acceptor wavelength and double-acceptor labeled proteins will not be excited. Moreover, the distance between fluorophores should be the same, regardless of which cysteine site a given fluorophore attaches to, especially when using the isotropic lanthanides as a donor, so the need to specify a given site to receive either the donor or acceptor is unnecessary. Intensity may be affected with a heterogeneous population, but should still be more than sufficient to be detected.
When planning experiments, the choice of fluorophores should be dictated by the R0 value of the pair as well as the expected distance range being measured. The R0 value is defined by the following equation:
<img alt="Equation 3" fo:content-width="2.5in" src="/files/ftp_upload/51895/51895eq3.jpg" width="250" />&#160;Eq. 3
where, R0&#160;is the F&#246;rster radius in Angstroms, &#954;2 is the orientation factor between the two dyes (usually assumed to be 2/3), &#981;D is the quantum yield of the donor, J is the spectral overlap integral between the donor&#39;s emission spectrum and the acceptor&#39;s absorbance spectrum in M-1cm-1nm4, and n is the refractive index of the medium1.
Our laboratory has added a modification to the conventional LRET technique by introducing a protease recognition site between the donor and acceptor label sites on the protein being probed. This modification allows for investigation in non-purified systems such as whole mammalian cells7. This technique is particularly useful when using cysteines as sites for labeling, since in the process of labeling with maleimide-conjugated dyes that bind to cysteine sulfhydryl groups, other proteins on the cells that have cysteines are also labeled. However, by including protease cleavage sites on the protein of interest and measuring lifetimes before and after cleavage, the experimenter can quantitatively subtract the background signal after protease cleavage from the raw signal. This subtraction isolates the specific signal arising from the protein of interest (Figure 2). Using the modification described above, LRET can be used to measure distance changes between the terbium chelate donor and the acceptor probe on a protein, and thus monitor conformational changes in the protein&#8217;s near physiological state without the requirement for purification.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51895/51895fig1highres.jpg" width="500" /> 
Figure 1.The absorption and emission spectra of chelated terbium in black, as well as a representative acceptor, Alexa 488, in red. Notice the multiple emission peaks and the sharp, narrow emission range for each peak of terbium chelate. This pattern allows for terbium to be used with a variety of acceptor fluorophores and facilitates the measurement of sensitized emission within those ranges where terbium shows no emission. Terbium&#8217;s emission peak at 486 nm overlaps quite well with the absorption peak of Alexa 488, allowing for resonance energy transfer to occur between the two fluorophores. A wavelength of 515 nm is an excellent choice to detect sensitized emission for this pair as it is in the valley between the terbium emission peaks, and quite near Alexa 488&#8217;s emission peak of 520 nm. Note that being near the acceptor peak, though desirable, is not required&#8212;565 nm is still able to detect Alexa 488 emission without also detecting terbium emission.
<table border="0" cellpadding="0" cellspacing="0" width="374">
	<tbody>
		<tr height="27">
			<td height="27" style="height:27px;width:147px;">Acceptor Fluorophore</td>
			<td style="width:45px;">R0 (&#197;)</td>
			<td style="width:183px;">Emission wavelength (nm)</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:147px;">Atto 465</td>
			<td align="right" style="width:45px;">36</td>
			<td align="right" style="width:183px;">508</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:147px;">Fluorescein</td>
			<td align="right" style="width:45px;">45</td>
			<td align="right" style="width:183px;">515</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:147px;">Alexa 488</td>
			<td align="right" style="width:45px;">46</td>
			<td align="right" style="width:183px;">515</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:147px;">Alexa 680</td>
			<td align="right" style="width:45px;">52</td>
			<td align="right" style="width:183px;">700</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:147px;">Alexa 594</td>
			<td align="right" style="width:45px;">53</td>
			<td align="right" style="width:183px;">630</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:147px;">Alexa 555</td>
			<td align="right" style="width:45px;">65</td>
			<td align="right" style="width:183px;">565</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:147px;">Cy3</td>
			<td align="right" style="width:45px;">65</td>
			<td align="right" style="width:183px;">575</td>
		</tr>
	</tbody>
</table>
Table 1. A list of commonly used acceptor fluorophores for LRET using terbium chelate as the donor11. The R0 values were measured when the donor and acceptor were attached to the soluble agonist binding domain of AMPA receptors. It is ideal to measure the R0 value again for each new system being studied.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51895/51895fig2highres.jpg" width="500" /> 
Figure 2. An overview of the LRET method presented. (A) The AMPA receptor is a membrane protein which undergoes conformational changes upon ligand-binding. The clamshell-shaped ligand-binding domain is circled here in red. (B) The ligand-binding domain of AMPA when not bound to protein exists in an open conformation (left). When bound to ligand glutamate, the protein closes around its ligand (right). By placing fluorophores at probative sites on the LBD, the nature of this conformational change can be seen as the distance between the fluorophores changes, which will then affect fluorescence lifetime. (C) When labeling whole cells, labeling of both the protein of interest as well as background membrane proteins may occur (left). After protease cleavage, LRET signal from the protein of interest will disappear due to the release of a soluble fragment, leaving background signal intact (right). This background signal can then be subtracted from the raw signal.

<table><tbody><tr><td>LanthaScreen Thiol Reactive Tb Chelate</td><td>Life Technologies</td><td>PV3579</td><td></td></tr><tr><td>Acceptor fluorophore-Fluorescein-5-Maleimide</td><td>Life Technologies&amp;nbsp;</td><td>F-150</td><td>Choice of acceptor depends on the specific experiment</td></tr><tr><td>Acceptor fluorophore-Alexa Fluor 488 C5 Maleimide</td><td>Life Technologies&amp;nbsp;</td><td>A-10254</td><td>Choice of acceptor depends on the specific experiment</td></tr><tr><td>Acceptor fluorophore-Alexa Fluor 555 C2 Maleimide</td><td>Life Technologies&amp;nbsp;</td><td>A-20346</td><td>Choice of acceptor depends on the specific experiment</td></tr><tr><td>Acceptor fluorophore-Alexa Fluor 594 C5 Maleimide</td><td>Life Technologies&amp;nbsp;</td><td>A-10256</td><td>Choice of acceptor depends on the specific experiment</td></tr><tr><td>Acceptor fluorophore-Alexa Fluor 680 C2 Maleimide</td><td>Life Technologies&amp;nbsp;</td><td>A-20344</td><td>Choice of acceptor depends on the specific experiment</td></tr><tr><td>QuantaMaster 3-SS</td><td>Photon Technology International</td><td></td><td>Spectrofluorometer should have pulsed excitation with the ability to measure lifetimes in the millisecond range</td></tr><tr><td>FluoreScan 2.0</td><td>Photon Technology International</td><td></td><td>Data Acquisition Software used in manuscript. Software provided with fluorescence instrument</td></tr><tr><td>Origin 8.6</td><td>OriginLab</td><td></td><td>Origin is the data analysis software used in protocol. Can use other similar data analysis software</td></tr><tr><td>Quartz cuvette</td><td>Starna Cells, Inc</td><td>3-Q-10</td><td></td></tr><tr><td>Stir bar&amp;nbsp;</td><td>Bel-Art Products</td><td>F37119-0007</td><td>Used in cuvette to keep cells in suspension. Can use any stir bar that fits the cuvette</td></tr></tbody></table>

luminescence resonance energy transfer to study conformational changes in membrane proteins expressed in mammalian cells

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 &#197;. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.

A successful LRET measurement with a lanthanide donor should have a donor only lifetime in the millisecond time range. The lifetime of sensitized LRET emission for the protein labeled with both the donor and the acceptor will be notably shorter, with a lifetime in the microsecond range after background subtraction Figure 3. Protease cleavage results in an increase in lifetime that should become stable over time (i.e. no longer changing), showing that the protease cleavage is complete Fig...

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Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range ...

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Research

JoVE Journal

Bioengineering

12.0K Views.  University of Texas Health Science Center at Houston. We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Video: Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

Biochemistry

Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately 20 nm to accommodate large tRNA substrates at the A-, P- and E-sites. Consequently, the ribosome dynamics are naturally de-phased quickly. Single molecule method can detect each ribosome separately and distinguish inhomogeneous populations, which is essential to reveal the complicated mechanisms of multi-component systems. We report the details of a smFRET method based on the Nikon Ti2 inverted microscope to probe the ribosome dynamics between the ribosomal protein L27 and tRNAs. The L27 is labeled at its unique Cys 53 position and reconstituted into a ribosome that is engineered to lack L27. The tRNA is labeled at its elbow region. As the tRNA moves to different locations inside the ribosome during the elongation cycle, such as pre- and post- translocation, the FRET efficiencies and dynamics exhibit differences, which have suggested multiple subpopulations. These subpopulations are not detectable by ensemble methods. The TIRF-based smFRET microscope is built on a manual or motorized inverted microscope, with home-built laser illumination. The ribosome samples are purified by ultracentrifugation, loaded into a home-built multi-channel sample cell and then illuminated via an evanescent laser field. The reflection laser spot can be used to achieve feedback control of perfect focus. The fluorescence signals are separated by a motorized filter-turret and collected by two digital CMOS cameras. The intensities are retrieved via the NIS-Elements software.

Single molecule fluorescence energy transfer is a method that tracks the tRNA dynamics during ribosomal protein synthesis. By tracking individual ribosomes, inhomogeneous populations are identified, which shed light on mechanisms. This method can be used to track biological conformational changes in general to reveal dynamic-function relationships in many other complexed biosystems. Single molecule methods can observe non-rate limiting steps and low-populated key intermediates, which are not accessible by conventional ensemble methods due to the average effect.

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately ...

Dual-Color Fluorescence Cross-Correlation Spectroscopy to Study Protein-Protein Interaction and Protein Dynamics in Live Cells

We present a protocol and workflow to perform live cell dual-color fluorescence cross-correlation spectroscopy (FCCS) combined with F&#246;rster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques. In dual-color FCCS, where the fluctuations in fluorescence intensity represent the dynamic &#34;fingerprint&#34; of the respective fluorescent biomolecule, we can probe co-diffusion or binding of the receptors. FRET, with its high sensitivity to molecular distances, serves as a well-known &#34;nanoruler&#34; to monitor intramolecular changes. Taken together, conformational changes and key parameters such as local receptor concentrations and mobility constants become accessible in cellular settings.
Quantitative fluorescence approaches are challenging in cells due to high noise levels and the vulnerability of the sample. Here we show how to perform this experiment, including the calibration steps using dual-color labeled &#946;2-adrenergic receptor (&#946;2AR) labeled with eGFP and SNAP-tag-TAMRA. A step-by-step data analysis procedure is provided using open-source software and templates that are easy to customize.
Our guideline enables researchers to unravel molecular interactions of biomolecules in live cells in situ with high reliability despite the limited signal-to-noise levels in live cell experiments. The operational window of FRET and particularly FCCS at low concentrations allows quantitative analysis at near-physiological conditions.

We present an experimental protocol and data analysis workflow to perform live cell dual-color fluorescence cross correlation spectroscopy (FCCS) combined with F&#246;rster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques.

We present a protocol and workflow to perform live cell dual-color fluorescence cross-correlation spectroscopy (FCCS) combined with F&#246;rster Resonance ...

F&#246;rster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between biomolecules in vitro as well as within cells. In FRET, the efficiency of energy transfer, measured by changes in fluorescence intensity or lifetime, relates to the distance between two fluorescent molecules or labels. Determination of dynamics and conformational changes from the distances are just some examples of applications of this method to biological systems. Under certain conditions, this methodology can add to and enhance existing X-ray crystal structures by providing information regarding dynamics, flexibility, and adaptation to binding surfaces. We describe the use of FRET and associated distance determinations to elucidate structural properties, through the identification of a binding site or the orientations of dimer subunits. Through judicious choice of labeling sites, and often employment of multiple labeling strategies, we have successfully applied these mapping methods to determine global structural properties in a protein-DNA complex and the SecA-SecYEG protein translocation system. In the SecA-SecYEG system, we have used FRET mapping methods to identify the preprotein-binding site and determine the local conformation of the bound signal sequence region. This study outlines the steps for performing FRET mapping studies, including identification of appropriate labeling sites, discussion of possible labels including non-native amino acid residues, labeling procedures, how to perform measurements, and interpreting the data.

The study details the methodology of FRET mapping including the selection of labeling sites, choice of dyes, acquisition, and data analysis. This methodology is effective at determining binding sites, conformational changes, and dynamic motions in protein systems and is most useful if performed in conjunction with existing 3-D structural information.

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between ...

Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET

The ability of cells to respond to external signals is essential for cellular development, growth, and survival. To respond to a signal from the environment, a cell must be able to recognize and process it. This task mainly relies on the function of membrane receptors, whose role is to convert signals into the biochemical language of the cell. G protein-coupled receptors (GPCRs) constitute the largest family of membrane receptor proteins in humans. Among GPCRs, metabotropic glutamate receptors (mGluRs) are a unique subclass that function as obligate dimers and possess a large extracellular domain that contains the ligand-binding site. Recent advances in structural studies of mGluRs have improved the understanding of their activation process. However, the propagation of large-scale conformational changes through mGluRs during activation and modulation is poorly understood. Single-molecule fluorescence resonance energy transfer (smFRET) is a powerful technique to visualize and quantify the structural dynamics of biomolecules at the single-protein level. To visualize the dynamic process of mGluR2 activation, fluorescent conformational sensors based on unnatural amino acid (UAA) incorporation were developed that allowed site-specific protein labeling without perturbation of the native structure of receptors. The protocol described here explains how to perform these experiments, including the novel UAA labeling approach, sample preparation, and smFRET data acquisition and analysis. These strategies are generalizable and can be extended to investigate the conformational dynamics of a variety of membrane proteins.

This study presents a detailed procedure to perform single-molecule fluorescence resonance energy transfer (smFRET) experiments on G protein-coupled receptors (GPCRs) using site-specific labeling via unnatural amino acid (UAA) incorporation. The protocol provides a step-by-step guide for smFRET sample preparation, experiments, and data analysis.

The ability of cells to respond to external signals is essential for cellular development, growth, and survival. To respond to a signal from the ...

CRAF-14-3-3 Interaction Analysis in Live Cells Using a BRET-Based Approach

Bioluminescence Resonance Energy Transfer (BRET)-Based Assay for Measuring Interactions of CRAF with 14-3-3 Proteins in Live Cells

CRAF is a primary effector of RAS GTPases and plays a critical role in the tumorigenesis of several KRAS-driven cancers. In addition, CRAF is a hotspot for germline mutations, which are shown to cause the developmental RASopathy, Noonan syndrome. All RAF kinases contain multiple phosphorylation-dependent binding sites for 14-3-3 regulatory proteins. The differential binding of 14-3-3 to these sites plays essential roles in the formation of active RAF dimers at the plasma membrane under signaling conditions and in maintaining RAF autoinhibition under quiescent conditions. Understanding how these interactions are regulated and how they can be modulated is critical for identifying new therapeutic approaches that target RAF function. Here, I describe a bioluminescence resonance energy transfer (BRET)-based assay for measuring the interactions of CRAF with 14-3-3 proteins in live cells. Specifically, this assay measures the interactions of CRAF fused to a Nano luciferase donor and 14-3-3 fused to a Halo tag acceptor, where the interaction of RAF and 14-3-3 results in donor-to-acceptor energy transfer and the generation of the BRET signal. The protocol further shows that this signal can be disrupted by mutations shown to prevent 14-3-3 binding to each of its high-affinity RAF docking sites. This protocol describes the procedures for seeding, transfecting, and replating the cells, along with detailed instructions for reading BRET emissions, performing data analysis, and confirming protein expression levels. In addition, example assay results, along with optimization and troubleshooting steps, are provided.

Author Spotlight: Integrating BRET-Based Assays and Rare Mutation Analysis to Decipher RAF Kinase Regulation in Live Cells

This protocol describes a BRET-based assay for measuring the interactions of the CRAF kinase with 14-3-3 proteins in live cells. The protocol outlines steps for preparing the cells, reading BRET emissions, and data analysis. An example result with identification of appropriate controls and troubleshooting for assay optimization is also presented.

CRAF is a primary effector of RAS GTPases and plays a critical role in the tumorigenesis of several KRAS-driven cancers. In addition, CRAF is a hotspot ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Biology

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live cells. BRET is the non-radiative transfer of energy from a 'donor' luciferase enzyme to an 'acceptor' fluorescent protein. In the most common configuration of this assay, the donor is Renilla reniformis luciferase and the acceptor is Yellow Fluorescent Protein (YFP). Because the efficiency of energy transfer is strongly distance-dependent, observation of the BRET phenomenon requires that the donor and acceptor be in close proximity. To test for an interaction between two proteins of interest in cultured mammalian cells, one protein is expressed as a fusion with luciferase and the second as a fusion with YFP. An interaction between the two proteins of interest may bring the donor and acceptor sufficiently close for energy transfer to occur. Compared to other techniques for investigating protein-protein interactions, the BRET assay is sensitive, requires little hands-on time and few reagents, and is able to detect interactions which are weak, transient, or dependent on the biochemical environment found within a live cell. It is therefore an ideal approach for confirming putative interactions suggested by yeast two-hybrid or mass spectrometry proteomics studies, and in addition it is well-suited for mapping interacting regions, assessing the effect of post-translational modifications on protein-protein interactions, and evaluating the impact of mutations identified in patient DNA.

Interactions between proteins are fundamental to all cellular processes. Using Bioluminescence Resonance Energy Transfer, the interaction between a pair of proteins can be monitored in live cells and in real time. Furthermore, the effects of potentially pathogenic mutations can be assessed.

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live ...

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...

Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Summary

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Chapters in this video

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

Dual-Color Fluorescence Cross-Correlation Spectroscopy to Study Protein-Protein Interaction and Protein Dynamics in Live Cells

Förster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET

Bioluminescence Resonance Energy Transfer (BRET)-Based Assay for Measuring Interactions of CRAF with 14-3-3 Proteins in Live Cells

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking