Roche's Publication Grant covered the Free Access publication and production of this article. The author also thanks E. Premkumar Reddy, Richard V. Mettus, Stephen C. Cosenza, Sau Ying Yip and Sol D. Gloria for their support and critical reading of the manuscript.

1. Cloning of the tdTomato-tagged Wild-type and Motorless KIF5B Proteins
<ol>
	<li>Amplify the cDNAs for the human wild-type and motorless KIF5B proteins using the primers in Table 1, Taq DNA polymerase (5 units for 100 &#956;l), dNTP mix (2 mM for each deoxynucleotide) and its 10X buffer for 30 cycles. Each cycle consists of a denaturation step (95 &#176;C for 30 sec), an annealing step (45 &#176;C for 30 sec) and an extension step (72 &#176;C for 3 min).</li>
	<li>Extract the amplified DNA product with equal volume of phenol/ chloroform (1:1). 
	Note: Phenol is combustible and can cause burns. Chloroform is hazardous. Avoid direct contact with them and use them under a chemical fume hood. Alternatively, PCR products can be purified by various kits.
	<ol>
		<li>Spin at 18,000 x g in a microcentrifuge at room temperature for 1 min. Transfer the aqueous solution to a new tube and extract with an equal volume of chloroform.</li>
		<li>Spin at 18,000 x g in a microcentrifuge at room temperature for 0.5 min. Transfer the aqueous solution to a new tube.</li>
		<li>Mix the aqueous solution with one tenth volume of 3 M Na-acetate and two volumes of absolute ethanol.</li>
		<li>Spin at 18,000 x g in a microcentrifuge at room temperature for 5 min. Discard the aqueous solution.</li>
		<li>Wash the DNA pellet with two volume of 75% ethanol. Discard the aqueous solution and air dry the DNA pellet at room temperature for 5 min. Resuspend the DNA pellet in 34 &#956;l water.</li>
	</ol>
	</li>
	<li>Digest the amplified products in a final volume of 40 &#956;l with restriction enzymes SalI (10 units) and BamHI (10 units) and 4 &#956;l of their 10X buffer at 37 &#176;C for two hr. Resolve the digested products in an agarose gel (1.0% at 100 V) containing ethidium bromide (0.2 &#956;g/ml). 
	Note: Ethidium bromide is a potent mutagen and can be absorbed through skin. Therefore, it is important to avoid direct or indirect contact with ethidium bromide.</li>
	<li>Cut out the correct bands for purification by columns. Weigh the agarose gel containing the DNA fragment. Then dissolve it in the solubilization buffer (300 &#956;l for 0.1 g) at 37 &#176;C for about 20 min.</li>
	<li>Add the resulted solution to a column and spin in a microcentrifuge at room temperature for 5 sec. Discard the flow-through.</li>
	<li>Wash the column with 0.5 ml solubilization buffer by repeating the step 1.5. Wash the column again with 0.75 ml wash buffer by repeating the step 1.5. Spin the column for 2 min to get rid of the remaining wash buffer.</li>
	<li>Elute the DNA fragment with 50 &#956;l water by repeating step 1.5. Estimate the concentration of the DNA fragment by running an aliquot of it against a DNA size ladder in an agarose gel (1.0% at 100 V) containing ethidium bromide (0.2 &#956;g/ml).</li>
	<li>Ligate the purified products (about 100 ng) with the ptdTomato-C1 vector17 (about 100 ng) previously digested with the same restriction enzymes in 10 &#956;l using T4 DNA ligase (2,000 units) and 1 &#956;l 10X ligation buffer at room temperature for 16 hr. 
	Note: The wild-type and motorless mutant contains amino acids from 2 to 963 and 584 to 963, respectively. Previously, a larger motorless KIF5B mutant was used to identify its function9. A smaller motorless KIF5B mutant was used for the present studies to increase its expression levels.</li>
</ol>
2. Immunofluorescence Microscopy
<ol>
	<li>For live-cell or indirect fluorescence imaging with magnification at or below 40X, seed 0.2-0.3 million HeLa cells in 1 ml complete medium [Dulbecco&#39;s Modified Eagle Medium (DMEM) with 10% fetal bovine serum (FBS)] in each well of a six-well plate. Grow at 37 &#176;C with 5 % CO2.</li>
	<li>For indirect immunofluorescence studies with magnifications above 40X, wash cover glasses (18 mm x 18 mm; 1.5 thickness) in absolute ethanol briefly and air-dry them in the wells of a six-well plate, inside a biosafety cabinet to avoid contamination.
	<ol>
		<li>Seed 0.2-0.3 million cells in 1 ml complete DMEM medium in each well of a six-well plate.</li>
	</ol>
	</li>
	<li>Preparation of Transfection Complex
	<ol>
		<li>The next day, inside a biosafety cabinet, dilute 0.6 &#181;g (total) of the expression plasmid for tdTomato-tagged wild-type or motorless KIF5B in the presence or absence of an expression plasmid for a cargo candidate protein (pTagCFP-c-MYC5) with 0.1 ml transfection medium in a 1.6 ml microcentrifuge tube. 
		Note: The vector pTagCFP-c-MYC expresses the TagCFP-tagged c-MYC.</li>
		<li>Add 1.8 &#181;l of the transfection reagent to 0.1 ml transfection medium in another 1.6 ml microcentrifuge tube. Typically, prepare a master mix of diluted transfection reagent enough for 12 samples.</li>
		<li>Incubate for 5 min, at room temperature.</li>
		<li>Add the diluted 0.1 ml transfection reagent to the diluted 0.1 ml DNAs. Mix the contents by inverting the tubes. Spin the tubes for 5 sec in a microcentrifuge.</li>
		<li>Incubate for 45 min, at room temperature.</li>
	</ol>
	</li>
	<li>Washing of Cells
	<ol>
		<li>Wash the seeded cells three times with phosphate buffered saline (PBS) during the incubation period for the formation of the transfection complex.</li>
		<li>Replace the medium of the seeded cells in each well with 0.8 ml pre-warmed transfection medium. Return the plates to the incubator at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Transfection of Cells
	<ol>
		<li>After incubation of 45 min, add 0.2 ml of the DNA/transfection reagent complex (prepared in step 2.3) dropwise to each well of the plate.</li>
		<li>Rock the 6 well plate gently for 5 sec, before they are returned to the incubator at 37 &#176;C.</li>
		<li>After 6-8 hr, replace the medium with the complete DMEM medium.</li>
	</ol>
	</li>
	<li>For Live-cell Fluorescence Imaging18
	<ol>
		<li>After an additional 16 hr of incubation at 37 &#176;C, add the DNA intercalating stain Hoechst 33342 to the medium to a final concentration of 1 &#181;M. 
		Note: Hoechst 33342 is a cell-permeable, DNA intercalating dye that stains the cell nuclei.</li>
		<li>Incubate the transfected cells for 10 min in the incubator at 37 &#176;C before examining them for the expression of fluorescent proteins by fluorescent microscopy using a 40X objective. The filter sets for DAPI, CFP, FITC and Cy3 are (Ex350 nm/ Em460 nm), (Ex436 nm/ Em480 nm), (Ex470 nm/ Em525 nm) and (Ex545 nm/ Em605 nm), respectively. 
		Note: If the background is high due to auto-fluorescence of the complete DMEM medium, medium without phenol red is used.</li>
	</ol>
	</li>
</ol>
3. For Indirect Immunofluorescence Studies
<ol>
	<li>Preparation of Paraformaldehyde Solution
	<ol>
		<li>Weigh paraformaldehyde powder (4 g per 100 ml PBS) and add it to PBS. 
		Note: Paraformaldehyde is a flammable solid and a potential cancer hazard. It irritates eyes, respiratory system and skin. Therefore, it is important to avoid contact with or inhalation of paraformaldehyde.</li>
		<li>Add NaOH to the solution (150 &#181;l 10 N NaOH/100 ml).</li>
		<li>Keep the solution at 37 to 42 &#176;C for 2-3 hr with occasional shaking.</li>
		<li>Adjust pH to 7.0 by adding glacial acetic acid to the solution (about 75 &#181;l/ 100 ml) after the paraformaldehyde powder is dissolved.</li>
	</ol>
	</li>
	<li>Target Proteins Staining
	<ol>
		<li>After further incubation for 16 hr at 37 &#176;C, wash the transfected cells at room temperature once with PBS by swirling the six-well plate for 5 sec.</li>
		<li>Fix the cells with 1 ml of freshly prepared, 4% paraformaldehyde solution per well. Incubate at room temperature for 30 min.</li>
		<li>Wash the cells with PBS once by swirling for 5 sec the six-well plate. Discard the solution.</li>
		<li>Incubate the cells with 1 ml of 0.1% Triton-X 100 (in PBS) at room temperature for 30 min. The detergent Triton-X 100 will permeabilize the cell membrane to allow access of antibodies to their intracellular targets.</li>
		<li>Wash the cells with PBS four times by swirling the six-well plate for 5 sec each time. Discard the solution after each wash.</li>
		<li>Incubate the cells with 1 ml of primary antibodies (c-MYC rabbit antibody or p53 mouse antibody; 0.1 &#181;g/ml) in a 10% FBS in PBS solution at room temperature by rocking for 4 hr.</li>
		<li>Wash with PBS four times by swirling the six-well plate for 5 sec each time. Discard the solution after each wash.</li>
		<li>Incubate with 1 ml of fluorescent dye-conjugated secondary antibodies (Alexa Fluor 488-conjugated anti-rabbit or anti-mouse IgG antibody; 0.5 &#181;g/ml) in 10% FBS in PBS at room temperature in the dark by rocking for 2 hr.</li>
		<li>Wash with PBS four times by swirling the six-well plate for 5 sec each time. Discard the solution after each wash.</li>
	</ol>
	</li>
	<li>Nuclear Staining and Mounting
	<ol>
		<li>Incubate the cells with the 1 ml of DNA intercalating dye 4&#39;,6-diamidino-2-phenylindole (DAPI; 0.5 &#181;g/ml) in PBS at room temperature, in dark for 10 min. Proceed to step 3.4 when no cover glasses are utilized. 
		Note: DAPI is a DNA intercalating dye that stains the cell nuclei.</li>
		<li>Apply 10 &#956;l of mounting solution onto each microscope slide. Mount each cover glass over the mounting solution on a microscope slide.</li>
		<li>Incubate at room temperature in the dark overnight.</li>
		<li>Seal the edges of the cover glasses with nail polish. Place in the dark inside a fume hood overnight to remove the nail polish fumes.</li>
	</ol>
	</li>
	<li>Fluorescence Microscopy19
	<ol>
		<li>Next day, examine the cells by fluorescent microscopy using a 40X objective. The filter sets for DAPI, CFP, FITC and Cy3 are (Ex350 nm/ Em460 nm), (Ex436 nm/ Em480 nm), (Ex470 nm/ Em525 nm) and (Ex545 nm/ Em605 nm), respectively.</li>
	</ol>
	</li>
</ol>

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Free Access publication and production of this article is paid for by Roche.

The method described utilizes the properties of the motorless KIF5B mutant, which lacks the ability to move along the microtubules, but retains the ability to form dimers with the wild-type KIF5B and, thereby, allow the tetrameric protein to interact with the same cargo proteins as the wild-type KIF5B. Motorless KIF5B, therefore, acts as a dominant negative mutant and forms mislocalized aggregates with its cargos. This method is proven to identify the Kinesin-1 cargo c-MYC (Figure 1)5. In this article, the same motorless KIF5B mutant was used to identify p53 as another potential cargo of Kinesin-1 (Figure 2). This shows that the methodology is feasible to identify other cargos of Kinesin-1. Furthermore, the specificity of the mutant is provided by the lack of effect of the mutant on the negative control protein c-Fos (Figure 3).
In this protocol, the tdTomato-tagged wild-type or motorless KIF5B protein is coexpressed with another fluorescent protein-tagged candidate cargo protein. In this case, live-cell fluorescence microscopy and imaging are performed. The formation of the aggregates can be traced by time-lapse imaging. Alternatively, the tdTomato-tagged protein is expressed alone and the candidate cargo protein at its physiological levels is visualized by indirect immunofluorescence microscopy using specific antibodies. The fluorescent protein tdTomato is chosen for its brightness and photostability17. If the background is high due to auto-fluorescence of the complete DMEM medium, medium without phenol red is used.
The motorless KIF5B mutant forms dimers with the wild-type KIF5B stoichiometrically. Therefore, it is critical to express sufficient amount of motorless KIF5B mutant to form dimers with the wild-type counterpart to inhibit its function and to form aggregates. To address this issue, optimization of expression of the motorless mutant is essential. It is achieved by using a small motorless mutant containing the dimerization domain. In addition, optimization of the appropriate transfection reagent and protocol is also necessary. The duration of incubation of HeLa cells with the DNA/ transfection reagent was optimized in HeLa cells for expression of exogenous proteins and cell viability. The incubation duration may have to be optimized for other cell lines.
For magnifications between 4X to 40X, no cover glasses are required. Cells can be directly examined in the wells. Therefore, in this case, the protocol is inexpensive and convenient. For magnifications above 40X, cells are grown on cover glasses in the wells and, after staining, are mounted on microscope slides for examination under oil-immersion objectives.
Observation of aggregates of the motorless KIF5B protein is limited by the size of the cytoplasm. It is easy to detect the aggregates in many mammalian cells. However, it is relatively difficult to observe the aggregates in neuronal cells when the volume of the cytoplasm is small around the nuclei and in neurites.
The protocol is used to show the association between KIF5B and its cargos in addition to the in vivo yeast two-hybrid and biochemical pull-down assays13-16. All these assays determine the association under different physical conditions and their results can complement each other. Moreover, the advantage of this fluorescence assay over other assays is that it can show the regulation of subcellular localization of the cargos by KIF5B (Figures 1 and 2).
The protocol is not limited to KIF5B and can be used to identify cargos of other motor proteins, such as some other kinesin motors3 and some of the myosin motors23,24 used in intracellular transport of cargos3. These motor proteins also contain motor domains for movement along microtubules for kinesin motors and microfilaments for myosin motors, and coiled-coil segments for oligomerization. Most of them form homodimers3,23. Therefore, a similar strategy can be applied to them by creating their motorless dominant negative mutants to identify their cargos.

Kinesin-1 is a motor protein that mediates anterograde transport of its cargos1,2. It is a heterotetramer of the two subunits of Kinesin Light Chain 1 (KLC1) and two subunits of Kinesin Heavy Chains (KHCs). KIF5B1, a KHC, contains a motor domain at its N-terminal, which hydrolyzes ATP and converts the chemical energy to mechanical energy for movement along microtubules. Its C-terminal region contains the dimerization domain that interacts with KLC1, which associates with cargos. Kinesin-1 transports cargos such as vesicles, organelles and mRNAs3,4. Recently, KIF5B was shown to transport the nuclear transcription factor c-MYC for proteosomal degradation in the cytoplasm5. Three methodologies (chemical inhibitor, siRNA/ shRNA and dominant negative mutant) were used to inhibit Kinesin-1 function. They all induced aggregation of c-MYC in the cytoplasm. For the last methodology, c-MYC was only affected by the dominant negative mutant of KIF5B, but not by that of another related KIF5A motor protein, suggesting that the mutant does not exert general effects on the intracellular components (like microtubule disruption) or on protein aggregation. The dominant negative mutant of KIF5B also did not affect another transcription factor, suggesting that it does not exert general effects on transcription factors. Rather, it suggests that the dominant negative mutant exerts specific effects on its cargos.
The use of dominant negative mutants is common in the field of motor proteins. Similar motorless mutants of kinesins and myosins were used previously. They were mainly used to demonstrate the effects of the mutants on the subcellular localizations of their cargos or on cellular functions6-12. Less emphasis was put on the spatial relationship between the mutants and the cargos affected by them. However, in some incidences, the mutants were observed to co-localize with their cargos6,10.
The interaction between KIF5B and its associated proteins was previously confirmed by the in vivo yeast two-hybrid assay and biochemical pull-down assays such as co-immunoprecipitation and in vitro pull-down assays13-16. In this article, an additional visual method using fluorescence microscopy is described to identify KIF5B cargo proteins. The method makes use of a motorless KIF5B mutant that acts as a dominant negative mutant. It aggregates in the cytoplasm and induces aggregation of its cargos.
The tagging of wild-type and motorless KIF5B mutant with the fluorescent protein tdTomato17 enables their visualization by fluorescence microscopy. The tagged KIF5B proteins can be co-expressed with a candidate protein fused to a different fluorescent protein with spectral properties suitably separated from the KIF5B tag. The tagged proteins are observed directly in live cells under fluorescence microscopy. Induction of aggregation of the candidate protein by the motorless KIF5B mutant will confirm that the candidate protein is an in vivo cargo of KIF5B. Furthermore, the tdTomato-tagged KIF5B proteins can be expressed alone in the cells to study their effects on the endogenous cargo proteins. Later, immunofluorescence microscopy is conducted in which the transfected cells are fixed and stained with a specific antibody against the endogenous candidate protein, followed by an appropriate secondary antibody conjugated with a fluorescent dye. In this case, the endogenous candidate protein at its physiological level is studied. Similar motorless mutants of other motor proteins can be prepared to identify their cargos.

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			<td height="21" style="height:21px;width:296px;">absolute ethanol (200 proof)</td>
			<td style="width:123px;">Fisher Scientific</td>
			<td style="width:113px;">BP2818</td>
			<td style="width:88px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DAPI</td>
			<td>Sigma-Aldrich</td>
			<td>D9542</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hoechst 33342</td>
			<td>Sigma-Aldrich</td>
			<td>B2261</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Opti-MEM-I (transfection medium)</td>
			<td>Life Technologies</td>
			<td align="right">51985</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ProLong Diamond Antifade Mountant</td>
			<td>Life Technologies</td>
			<td>P36961</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">formaldehyde, para</td>
			<td>Fisher Scientific</td>
			<td>O4042-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triton-X100</td>
			<td>Fisher Scientific</td>
			<td>BP337-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">X-tremeGENE 9 DNA Transfection Reagent</td>
			<td>Roche</td>
			<td align="right">6365787001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Taq DNA polymerase</td>
			<td>Life Technologies</td>
			<td align="right">10342020</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PCR Grade Nucleotide Mix (dNTP Mix)</td>
			<td>Roche</td>
			<td align="right">12111424</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Microscope cover glass</td>
			<td>Fisher Scientific</td>
			<td>12-541A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">GeneRuler 1 kb Plus DNA ladder</td>
			<td>Life Technologies</td>
			<td>SM1333</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PureLink Quick Gel Extraction kit</td>
			<td>Life Technologies</td>
			<td>K210012</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">BamHI</td>
			<td>New England Biolab</td>
			<td>R0136</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">SalI</td>
			<td>New England Biolab</td>
			<td>R0138</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">T4 DNA ligase&#160;</td>
			<td>New England Biolab</td>
			<td>M0202T</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ethidium bromide</td>
			<td>Thermo Scientific</td>
			<td align="right">17898</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DMEM</td>
			<td>Life Technologies</td>
			<td>11995-065</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">c-MYC rabbit antibody</td>
			<td>Cell Signaling</td>
			<td align="right">5606</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">p53 mouse antibody</td>
			<td>Santa Cruz</td>
			<td>sc-126</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 488-conjugated anti-rabbit IgG antibody</td>
			<td>Life Technologies</td>
			<td>A11008</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 488-conjugated anti-mouse IgG antibody</td>
			<td>Life Technologies</td>
			<td>A11059</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">fluorescent microscope&#160;</td>
			<td colspan="2">Olympus IX71_Fluoview</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">computer software for imaging</td>
			<td>&#160;cellSens&#160;</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

identification of kinesin-1 cargos using fluorescence microscopy

Fluorescence microscopy is employed to identify Kinesin-1 cargos. Recently, the heavy chain of Kinesin-1 (KIF5B) was shown to transport the nuclear transcription factor c-MYC for proteosomal degradation in the cytoplasm. The method described here involves the study of a motorless KIF5B mutant for fluorescence microscopy. The wild-type and motorless KIF5B proteins are tagged with the fluorescent protein tdTomato. The wild-type tdTomato-KIF5B appears homogenously in the cytoplasm, while the motorless tdTomato-KIF5B mutant forms aggregates in the cytoplasm. Aggregation of the motorless KIF5B mutant induces aggregation of its cargo c-MYC in the cytoplasm. Hence, this method provides a visual means to identify the cargos of Kinesin-1. A similar strategy can be utilized to identify cargos of other motor proteins.

Exogenously expressed wild-type KIF5B appeared homogeneously in the cytoplasm, while c-MYC appeared mainly in the nucleus (Figure 1A). However, the motorless KIF5B mutant formed aggregates in the cytoplasm (Figure 1A). Aggregation of the motorless KIF5B induced the aggregation of c-MYC. The percentage of cells expressing aggregated TagCFP-tagged c-MYC in the cells expressing wild-type KIF5B was low. However, it was significantly higher when the motorless KIF5B was co-expressed (Figure 1A). The observed co-localization of mutant KIF5B with c-MYC (Figure 1) suggests that KIF5B regulates the subcellular localization of c-MYC and c-MYC is a cargo of Kinesin-1. Negative controls in which the constructs were expressed alone are included in the lower panel of Figure 1A. The results show that there was no significant bleed-through of fluorescence emission. The motorless KIF5B also induced aggregation of the endogenous c-MYC (Figure 1B) and the transcription factor p53 (Figure 2), indicating that c-MYC and p53 are both the cargos of Kinesin-1 and KIF5B regulates the subcellular localization of both endogenous proteins. Together with the results that the Kinesin-1 inhibitor rose bengal lactone (RBL)20 induces formation of high molecular weight species for both c-MYC and p535, p53 is likely a cargo of Kinesin-1. It is interesting to note that the nuclear transcription factor p53, like c-MYC, is also exported from the nucleus to the cytoplasm for proteasomal degradation21,22. The movement of transcription factors by KIF5B appears to be specific because the expression of the motorless KIF5B mutant did not affect the subcellular localization c-Fos or cause it to aggregate (Figure 3). The above data demonstrate that the method employed in this publication allows for the high-specificity identification of Kinesin-1 cargos. A similar strategy may be applied to other motor proteins as well.
<img alt="Figure 1" src="/files/ftp_upload/53632/53632fig1.jpg" /> 
Figure 1: Expression of the motorless KIF5B mutant induces c-MYC aggregation in the cytoplasm. (A) HeLa cells were transfected with tdTomato-tagged wild-type (WT) KIF5B (red) and TagCFP-tagged c-MYC (blue). tdTomato-tagged WT KIF5B appeared mainly in the cytoplasm, while TagCFP-tagged c-MYC appeared mainly in the nucleus. However, the tdTomato-tagged motorless KIF5B mutant (red) formed filamentous or punctate aggregates in the cytoplasm. Expression of the motorless KIF5B induced the aggregation of c-MYC. The mutant KIF5B and c-MYC co-localized together (pink). Percentages of cells (%) indicating aggregates of CFP-c-MYC (%) are shown. The results are shown as mean&#177;SD (N = 3). Negative controls with expression of tdTomato-tagged KIF5B proteins or TagCFP-tagged c-MYC along with empty vectors (V) are shown in the lower panel. (B) Similar results were obtained with the endogenous c-MYC (green) when tdTomato-tagged WT or motorless KIF5B proteins (red) were expressed. The tdTomato-tagged motorless KIF5B mutant formed aggregates in the cytoplasm and induced aggregation of the endogenous c-MYC. Both kinds of aggregates co-localized together in the cytoplasm (orange). The nuclei were stained with the dye Hoechst 33342 or DAPI. Scale bar; 20 &#956;m. <a href="https://www.jove.com/files/ftp_upload/53632/53632fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53632/53632fig2.jpg" /> 
Figure 2: Expression of the motorless KIF5B mutant induces endogenous p53 aggregation in the cytoplasm. In HeLa cells, exogenous tdTomato-tagged wild-type (WT) KIF5B (red) appeared mainly in the cytoplasm, while the endogenous p53 (green) appeared mainly in the nucleus. In contrast, the tdTomato-tagged motorless KIF5B mutant (red) formed aggregates in the cytoplasm. Expression of the motorless KIF5B induced the aggregation of p53, resulting in the co-localization of KIF5B and p53 in the cytoplasm (yellow). The experiment was performed three times. The nuclei were stained with the dye DAPI. Scale bar; 20 &#956;m. <a href="https://www.jove.com/files/ftp_upload/53632/53632fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53632/53632fig3.jpg" /> 
Figure 3: Expression of the motorless KIF5B mutant does not induce aggregation of c-Fos in the cytoplasm. tdTomato-tagged wild-type (WT) KIF5B (red) appeared mainly in the cytoplasm of HeLa cells, while EGFP-c-Fos (green) appeared mainly in the nucleus. On the contrary, the tdTomato-tagged motorless KIF5B mutant (red) formed aggregates in the cytoplasm. Expression of the motorless KIF5B did not induce the aggregation of c-Fos. The experiment was performed four times. The nuclei of the cells were stained with the dye DAPI. Scale bar; 20 &#956;m. <a href="https://www.jove.com/files/ftp_upload/53632/53632fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>common reverse primer</td>
			<td>5&#8217;-AGAGGATCCTTACACTTGTTTGCCTCCTC-3&#8217;</td>
		</tr>
		<tr>
			<td>wild-type KIF5B forward primer</td>
			<td>5&#8217;- AGAGTCGACGCGGACCTGGCCGAGTGCAACATCAAAGT-3&#8217;</td>
		</tr>
		<tr>
			<td>motorless KIF5B forward primer</td>
			<td>5&#8217;- AGAGTCGACGATGAAGAGTTCACTGTTGC-3&#8217;</td>
		</tr>
	</tbody>
</table>
Table 1: Primer sequences for cloning the wild-type and motorless KIF5B proteins.

Watch this Scientific Journal Video about Identification of Kinesin-1 Cargos Using Fluorescence Microscopy at JoVE.com

Identification of Kinesin-1 Cargos Using Fluorescence Microscopy

Here, a protocol is presented to identify Kinesin-1 cargos. A motorless mutant of the Kinesin-1 heavy chain (KIF5B) aggregates in the cytoplasm and induces aggregation of its cargos. Both aggregates are detected under fluorescence microscopy. A similar strategy can be employed to identify cargos of other motor proteins.

Fluorescence microscopy is employed to identify Kinesin-1 cargos. Recently, the heavy chain of Kinesin-1 (KIF5B) was shown to transport the nuclear ...

identification-of-kinesin-1-cargos-using-fluorescence-microscopy

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7.8K Views.  Icahn School of Medicine at Mount Sinai. Here, a protocol is presented to identify Kinesin-1 cargos. A motorless mutant of the Kinesin-1 heavy chain (KIF5B) aggregates in the cytoplasm and induces aggregation of its cargos. Both aggregates are detected under fluorescence microscopy. A similar strategy can be employed to identify cargos of other motor proteins.

Video: Identification of Kinesin-1 Cargos Using Fluorescence Microscopy

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Fluorescence-microscopy Screening and Next-generation Sequencing: Useful Tools for the Identification of Genes Involved in Organelle Integrity

This protocol describes a fluorescence microscope-based screening of Arabidopsis seedlings and describes how to map recessive mutations that alter the subcellular distribution of a specific tagged fluorescent marker in the secretory pathway. Arabidopsis is a powerful biological model for genetic studies because of its genome size, generation time, and conservation of molecular mechanisms among kingdoms. The array genotyping as an approach to map the mutation in alternative to the traditional method based on molecular markers is advantageous because it is relatively faster and may allow the mapping of several mutants in a really short time frame. This method allows the identification of proteins that can influence the integrity of any organelle in plants. Here, as an example, we propose a screen to map genes important for the integrity of the endoplasmic reticulum (ER). Our approach, however, can be easily extended to other plant cell organelles (for example see1,2), and thus represents an important step toward understanding the molecular basis governing other subcellular structures.

A fundamental quest in cell biology is to define the mechanisms that underlie the identity of the organelles that make eukaryotic cells. Here we propose a method to identify the genes responsible for the morphological and functional integrity of plant organelles using fluorescence microscopy and next-generation sequencing tools.

This protocol describes a fluorescence microscope-based screening of Arabidopsis seedlings and describes how to map recessive mutations that alter the ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells possess several lines of defense against damage to biologically relevant molecules like nucleic acids, lipids and proteins. In addition to organelle dynamics (fusion/fission/motility/inheritance) and tightly controlled protease activity, the degradation of surplus, damaged or compromised organelles by autophagy (cellular &#8216;self-eating&#8217;) has received much attention from the scientific community. The regulation of autophagy is quite complex and depends on genetic and environmental factors, many of which have so far not been elucidated. Here a novel method is presented that allows the convenient study of autophagy in the filamentous fungus Penicillium chrysogenum. It is based on growth of the fungus on so-called &#8216;starvation pads&#8217; for stimulation of autophagy in a reproducible manner. Samples are directly assayed by microscopy and evaluated for autophagy induction / progress. The protocol presented here is not limited for use with P. chrysogenum and can be easily adapted for use in other filamentous fungi.

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

A convenient and powerful method for studying autophagy in Penicillium chrysogenum by using starvation pads (mixtures of agarose and tap water in a microscope slide containing a central cavity) is presented here.

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...