This work was supported by Grant 31003A-162643 by the Swiss National Science Foundation. We thank Nicole Beuret and the Biozentrum Imaging Core Facility (IMCF) for support.

1. Bacterial Transformation with Functionalized Nanobodies
NOTE: This protocol has been optimized for the expression, purification, and analysis of functionalized anti-GFP nanobodies as previously described30. Derivatization with other protein moieties might require modification of this standard protocol.
<ol>
	<li>Thaw chemocompetent bacteria (~100 &#181;L) suited for protein expression (e.g., Escherichia coli Rosetta BL21 (DE3) cells) by placing them on ice. 
	NOTE: Prepare chemocompetent bacterial cells according to standard lab procedures. Alternatively, chemically competent bacterial cells can be purchased commercially.</li>
	<li>Add 50 ng of a plasmid encoding functionalized nanobodies. To allow sufficient site-specific biotinylation of the nanobody reporter during bacterial expression, also add a threefold excess (150 ng) of a bacterial expression plasmid encoding biotin ligase BirA (see also Table 1 for plasmid information). Gently pipet up and down. 
	NOTE: If nanobody biotinylation is not necessary or the nanobody reporters are devoid of a biotin acceptor peptide (BAP), co-transformation with a plasmid encoding BirA is not required.</li>
	<li>Incubate the bacterial cells for 30 min on ice.</li>
	<li>Heat shock the bacterial cells by placing them for 1 min at 42 &#176;C in a water bath or a heating block.</li>
	<li>Add 1 mL of room-temperature (RT) Luria broth (LB) medium and incubate the transformed bacteria in a thermoshaker for 1 h at 37 &#176;C to allow phenotypic expression of resistance genes. To prepare 1 L LB medium, balance 5 g of yeast extract, 10 g of tryptone and 10 g of NaCl, fill up with water and sterilize by autoclaving. If the functionalized nanobody has been subcloned in an expression vector containing resistance to ampicillin/carbenicillin, step 1.5 can be omitted.</li>
	<li>Pellet the bacteria at 11,000 x g for 1 min and resuspend in 100 &#181;L of fresh LB medium.</li>
	<li>Plate the suspended bacteria on LB plates containing the respective antibiotics (e.g., with 50 &#181;g/mL kanamycin and 50 &#181;g/mL carbenicillin when the bacteria have been co-transformed with nanobody reporter and BirA as stated above (step 1.2)).</li>
	<li>Place the LB plates in an incubator at 37 &#176;C and let grow overnight (O/N). 
	NOTE: The protocol can be paused here by storing the LB plate with grown colonies at 4 &#176;C. Use parafilm to seal LB plates.</li>
</ol>
2. Bacterial Liquid Culture and Induction of Functionalized Nanobody Expression
<ol>
	<li>Pick a bacterial colony containing the vector of interest from the plate and let it grow in a flask containing 20 mL of LB medium supplemented with antibiotics O/N in a shaking incubator at 37 &#176;C (see also step 1.7 regarding selection antibiotics).</li>
	<li>Next day, dilute the grown 20 mL bacterial culture into a flask containing 1 L of LB medium with selection antibiotics.</li>
	<li>Continue to incubate the bacterial culture at 37 &#176;C until it reaches an OD600 of 0.6-0.7. Allow the culture to cool down to RT before inducing protein expression.</li>
	<li>Induce protein expression of functionalized nanobodies and BirA by the addition of 1 mL of 1 M isopropyl-&#946;-d-thiogalactopyranosid (IPTG) to a final concentration of 1 mM of the inducer (1:1,000 dilution). Also add 10 mL of a 20 mM d-biotin stock solution resulting in 200 &#181;M d-biotin in the growth medium to allow biotinylation of the BAP epitope present in functionalized nanobodies (see also Table 1 for plasmid information) 
	NOTE: The d-biotin stock solution is prepared in ddH2O and brought into solution by drop-wise addition of 500 mM NaH2PO4. Alternatively, d-biotin can also be dissolved in DMSO. To produce nanobodies fused to APEX2 (VHH-APEX2), the LB medium must be complemented in addition with 1 mM 5-aminolevulinic acid (dALA) hydrochloride to promote heme incorporation. dALA is prepared as a 100 mM stock solution in ddH2O.</li>
	<li>Incubate the IPTG-induced 1 L bacterial culture at 30 &#176;C for 4 h for VHH-2xTS, at 20 &#176;C or RT O/N for VHH-APEX2, or at 16 &#176;C O/N for VHH-mCherry (see also Table 1 for plasmid information). 
	NOTE: Expression conditions for a new nanobody construct must be optimized by the experimenter.</li>
	<li>Transfer the bacterial culture from the flask into a 1 L centrifugation bottle and pellet the cells at 5,000 x g at 4 &#176;C for 45 min. Decant the supernatant and continue with the purification. 
	NOTE: The protocol can be paused here by storing the bacterial pellet at -80 &#176;C indefinitely. The pellet for VHH-APEX2 or VHH-mCherry is typically brownish (because of the incorporated heme) or pink, respectively.</li>
</ol>
3. Purification of Functionalized Nanobodies
<ol>
	<li>If necessary, thaw the frozen bacterial pellet on ice (see also step 2.6).</li>
	<li>Add 30 mL of ice-cold binding buffer (20 mM imidazole in 1x PBS) to the bacterial cell pellet and resuspend by pipetting up and down. Transfer resuspended bacteria into a labeled 50 mL tube.</li>
	<li>Supplement the 30 mL binding buffer with 200 &#181;g/mL lysozyme, 20 &#181;g/mL DNase I, 1 mM MgCl2 and 1 mM phenylmethylsulfonyl fluoride (PMSF) and incubate first for 10 min at RT, and then for 1 h at 4 &#176;C on an end-over-end shaker.</li>
	<li>Mechanically lyse bacterial cells in a 50 mL tube by placing a tip sonicator into the suspension. Apply constant 3x 1 min pulses with 1 min cooling periods in-between.</li>
	<li>Centrifuge the bacterial lysate at 15,000 x g at 4 &#176;C for 45 min to pellet the debris and intact cells. Transfer the sonicated bacterial cell lysate either into an appropriate centrifugation bottle for centrifugation or divide the lysate into several 5 mL tubes for tabletop centrifugation.</li>
	<li>Transfer the supernatant into a new 50 mL tube and discard the pelleted debris.</li>
	<li>Store the cleared lysate on ice while preparing purification by immobilized metal affinity chromatography (IMAC). To isolate histidine-tagged functionalized nanobodies, employ single-use His columns (see Table of Materials) allowing fast and simple gravity-flow purification. 
	NOTE: Alternatively, batch purification instead of commercial columns can be also used.</li>
	<li>Mounting His Columns at a Metal Stand.
	<ol>
		<li>Taper off the columns&#8217; storage solution, and equilibrate the His column with 10 mL of binding buffer (20 mM imidazole in 1x PBS).</li>
		<li>Empty by gravity flow (the flow-through can be discarded).</li>
	</ol>
	</li>
	<li>Gradually load the cleared bacterial lysate (~30 mL) onto the column. Empty by gravity flow (the flow-through can be discarded).</li>
	<li>Wash the His column 2x with 10 mL of binding buffer (20 mM imidazole in 1x PBS).</li>
	<li>Elute the nanobodies with 2 mL of elution buffer (500 mM imidazole in 1x PBS) into a 2 mL tube and apply a buffer exchange.</li>
	<li>Buffer Exchange.
	<ol>
		<li>Equilibrate a desalting column (see Table of Materials) placed in a 50 mL tube column adapter 5 times with 5 mL of 1x PBS.</li>
		<li>Allow the buffer to enter the packed bed. Discard the flow-through.</li>
		<li>After the fifth PBS equilibration, spin the column at 1,000 x g for 2 min.</li>
		<li>Discard the flow-through.</li>
		<li>Place the column with the adapter onto a new 50 mL tube. Load the 2 mL of eluted functionalized nanobody (from step 3.11) onto the PBS-equilibrated desalting column and spin at 1,000 x g for 2 min and collect the eluate. 
		NOTE: Instead of commercial desalting columns, dialysis can be applied to exchange buffer composition.</li>
	</ol>
	</li>
	<li>Determine the protein (nanobody) concentration of the eluate using the bicinchoninic (BCA) or Bradford assay according to the manufacturer&#8217;s instructions. 
	NOTE: For nanobody uptake assays by GFP reporter cell lines, a stock concentration of 2-10 mg/mL is optimal.</li>
</ol>
4. Validation of Functionalized Nanobody Expression (Coomassie Staining)
<ol>
	<li>Prepare 10-12.5% gels for sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) according to standard protocols. 
	NOTE: Alternatively, use commercially available precast gradient gels.</li>
	<li>Pipet 20 &#181;g of purified functionalized nanobody into a 1.5 mL tube and boil in sample buffer at 95 &#176;C for 5 min. 
	NOTE: As an example, add 10 &#181;L of a 2 mg/mL nanobody stock solution into a 1.5 mL tube with 10 &#181;L of 2x sample buffer. If the volume is too small, further dilute nanobody in PBS before adding sample buffer.</li>
	<li>Load the purified nanobody boiled in sample buffer onto an SDS-polyacrylamide gel, and run according to the standard PAGE protocol until the reference dye (e.g., bromophenol blue) has reached the end of the separating gel, followed by Coomassie staining and destaining.</li>
	<li>Coomassie Gel Staining and Destaining.
	<ol>
		<li>Uncast the gel and stain it with Coomassie staining solution (5% of a 10 g/L Coomassie stock solution in 10% acetic acid and 45% methanol in ddH2O) for 20-30 min at RT on a moving shaking platform. The gel should be completely covered in the staining solution.</li>
		<li>Destain the gel 2-3 times with destain solution (7.5% acetic acid and 15% methanol in ddH2O) for 1 h each at RT, before leaving the gel O/N for further destaining. 
		NOTE: Excess Coomassie can be efficiently soaked up by placing kitchen paper towels around the gel.</li>
	</ol>
	</li>
	<li>Image the gel with an imager or camera of choice. 
	NOTE: Nanobody expression can be further validated by immunoblot analysis using antibodies targeting its epitopes (see also Figure 1A).</li>
</ol>
5. Uptake of Functionalized Nanobodies by Cultured Cells for Fluorescence Staining
<ol>
	<li>In a cell culture hood, seed ~400,000 to 500,000 HeLa cells stably expressing a GFP-tagged reporter protein on 18-mm glass coverslips (No. 1.5H) in 35-mm dishes or 6-well clusters with complete medium containing antibiotics (Dulbecco&#8217;s Modified Eagle medium, DMEM, supplemented with 10% fetal calf serum (FCS), 100 units/mL streptomycin, 2 mM l-glutamine, and 1.5 &#181;g/mL puromycin). 
	NOTE: Here, we use HeLa cells stably expressing EGFP-Calnexin, EGFP-CDMPR and TfR-EGFP for illustration purposes. Apart from stable cell lines, also transiently transfected HeLa cells can be used. Stable cell lines can be co-cultivated at this step with parental non-transduced HeLa cells, for direct comparison.</li>
	<li>Incubate the cells O/N at 37 &#176;C in a 7.5% CO2 incubator to proliferate. 
	NOTE: Cells should have a confluency of ~80% the next day.</li>
	<li>Pipet 2 &#181;L of a 2 mg/mL nanobody stock solution into a 15 or 50 mL tube containing 2 mL of complete medium (this volume is sufficient to cover a 35 mm dish or one well of a 6-well cluster, adjust the volume of medium and nanobody proportionally to include more cell culture dishes or clusters). 
	NOTE: For the purpose of illustration, we here use VHH-mCherry, since no further antibody staining is required to see nanobody internalized by the EGFP reporters (see Figure 2C).</li>
	<li>Remove growth medium from the cells and add 2 mL of prewarmed complete medium containing 2 &#181;g/mL functionalized nanobody per 35 mm dish or 6-well unit.</li>
	<li>Incubate the cells for 1 h at 37 &#176;C in a 7.5% CO2 incubator to allow reporter-mediated nanobody uptake. 
	NOTE: Depending on the planned experiment, the time of nanobody uptake needs to be adjusted. For the experiment shown here, 1 h is sufficient to reach steady-state of nanobody uptake by EGFP-CDMPR and TfR-EGFP.</li>
	<li>Remove the medium and wash the cells 2-3 times with 1 mL of 1x PBS at RT.</li>
	<li>Fix the cells with 1 mL of 3% paraformaldehyde (PFA) for 10 min at RT.</li>
	<li>Remove PFA solution and quench remaining fixative with 1 mL of 50 mM NH4Cl in 1x PBS for 5 min at RT. 
	NOTE: 3% PFA should be disposed of separately as fixative waste.</li>
	<li>Wash the cells 3 times with 1 mL of 1x PBS at RT.</li>
	<li>Permeabilize the cells with 1 mL of 0.1% Triton X-100 in 1x PBS for 10 min at RT. 
	NOTE: Although no permeabilization is required, EGFP and mCherry fluorescence is better when afterwards the cells are embedded in mounting medium.</li>
	<li>Wash the cells 3 times with 1 mL of 1x PBS at RT.</li>
	<li>Place the coverslips with forceps on a drop (~100 &#181;L) of 1% BSA in 1x PBS containing 5 &#181;g/mL DAPI (4&#39;,6-diamidino-2-phenylindole) for 5 min at RT.</li>
	<li>Place the coverslips back into the dish/well and wash 3 times with 1 mL of 1x PBS at RT.</li>
	<li>Label the glass slides and mount the cells with Fluoromount-G. Let the mounting medium harden in the dark for 3-4 h and store the glass slides at 4 &#176;C under light protection.</li>
	<li>Image staining patterns using a microscope of choice (e.g., Point Scanning Confocal upright microscope).</li>
</ol>
6. Uptake of Functionalized Nanobodies by Cultured Cells (for Sulfation Analysis)
<ol>
	<li>In a cell culture hood, seed ~400,000 to 500,000 HeLa cells stably expressing a GFP-tagged reporter protein in 35 mm dishes or on 6-well clusters with complete medium containing antibiotics (Dulbecco&#8217;s Modified Eagle medium, DMEM, supplemented with 10% fetal calf serum (FCS), 100 units/mL streptomycin, 2 mM l-glutamine, and 1.5 &#181;g/mL puromycin). 
	NOTE: As mentioned above, we here use HeLa cells stably expressing EGFP-Calnexin, EGFP-CDMPR and TfR-EGFP for illustration purposes. Apart from stable cell lines, also transiently transfected HeLa can be used.</li>
	<li>Incubate the cells O/N at 37 &#176;C in a 7.5% CO2 incubator to proliferate. 
	NOTE: Cells should have a confluency of ~80% next day.</li>
	<li>Remove the complete medium, wash 2 times with 1x PBS at RT, and starve the cells with sulfate-free medium (SFM) for 1 h at 37 &#176;C in a 7.5% CO2 incubator. 
	NOTE: SFM is prepared by adding 10 mL of MEM amino acids (50x) solution, 5 mL of L-glutamine (200 mM), 5 mL of vitamin solution (100x), and 900 &#181;L of CaCl2&#183;2H2O to 500 mL of Eagle&#8217;s balanced salts. Pass through a 0.22 &#181;m filter and aliquot.</li>
	<li>In a ventilated hood or bench designated for radioactivity work, prepare sulfate labeling medium containing 0.5 mCi/mL [35S]sulfate as sodium salt in SFM. 
	CAUTION: Carefully handle all solutions containing radioactivity. Only work in designated hoods or benches. All material (tips, tubes, plates, etc.) or wash buffers in contact with radioactivity have to be collected and disposed of separately. Do this for all steps below (step 6.5-6.20) as well.</li>
	<li>Add VHH-2xTS in 1x PBS into sulfate labeling medium to a final concentration of 2 &#181;g/mL. 
	NOTE: As control, also include VHH-std or another nanobody devoid of TS sites to demonstrate specific labeling.</li>
	<li>Replace SFM by 0.7 mL of sulfate labeling medium containing 0.5 mCi/mL [35S]sulfate and 2 &#181;g/mL VHH-2xTS and incubate the cells for 1 h at 37 &#176;C in a 7.5% CO2 incubator designated for work with radioactivity. 
	NOTE: Depending on the planned experiment, the time of nanobody sulfation needs to be adjusted. In addition, to determine TGN arrival kinetics, labeling is performed for different times (e.g., for 10 min, 20 min, 30 min, etc.).</li>
	<li>Remove the labeling medium and wash the cells 2-3 times with ice-cold 1x PBS on a cooling plate or ice.</li>
	<li>Add 1 mL of lysis buffer (PBS containing 1% Triton X-100 and 0.5 % deoxycholate) supplemented with 2 mM PMSF and 1x protease inhibitor cocktail and incubate the cells for 10-15 min on a rocking platform at 4 &#176;C.</li>
	<li>Scrape and transfer the lysate into a 1.5 mL tube, vortex the lysate, and place for 10-15 min on an end-over-end rotator at 4 &#176;C.</li>
	<li>Prepare a postnuclear lysate by centrifugation at 10,000 x g for 10 min at 4 &#176;C.</li>
	<li>Using a new 1.5 mL tube, prepare 20-30 &#181;L of a slurry of Nickel beads in a 1.5 mL tube and wash once with 1x PBS by gently pelleting them at 1,000 x g for 1 min. 
	NOTE: Instead of Nickel beads, streptavidin beads (if nanobody is biotinylated) or protein A beads with an IgG against an epitope of the nanobody (T7- or HA-tag) can be used.</li>
	<li>Transfer the postnuclear lysate into a 1.5 mL tube containing Nickel beads and incubate for 1 h at 4 &#176;C on an end-over-end rotator to isolate the nanobodies. Remove an aliquot (50-100 &#181;L) of the postnuclear lysate and boil in SDS sample buffer for subsequent immunoblot analysis of total cell-associated nanobody, GFP reporter and loading control.</li>
	<li>Wash the beads 3 times with 1x PBS or lysis buffer by gently pelleting at 1,000 x g for 1 min. 
	NOTE: For more stringent washing of beads, 20 mM imidazole can be included in PBS or lysis buffer.</li>
	<li>Carefully remove all washing buffer from the beads, add 50 &#181;L of 2x sample buffer, and boil for 5 min at 95 &#176;C.</li>
	<li>Load both boiled beads on a midi 12.5% SDS-PAGE gel and run according to the standard PAGE protocol until the reference dye (e.g., bromophenol blue) has reached the end of the separating gel. 
	NOTE: Immunoblot analysis of total cell-associated nanobody, GFP reporter and loading control can also be performed using a mini 12.5% SDS-PAGE or precast gradient gel.</li>
	<li>Fix the separating gel in ~30 mL of fixation buffer (10% acetic acid and 45% methanol in ddH2O) for 1 h at RT, wash the gel three times with ~50 mL of deionized water and prepare for gel drying.</li>
	<li>Drying SDS-PAGE Gels. 
	<ol>
		<li>Carefully place the fixed gel on stretched cling film and cover it with precut filter paper.</li>
		<li>Place the gel with the filter paper down on top of the drying platform, place the vacuum cover, and switch on the vacuum pump for gel drying. Dry the gel for 3-5 h.</li>
		<li>Remove the cling film and place the dried gel into an autoradiography cassette equipped with a Phosphor Screen.</li>
	</ol>
	</li>
	<li>Image autoradiograph using a Phosphor Screen Imager. Follow the instructions of the manufacturer. 
	NOTE: Depending on the strength of the signal, dried gels need to be exposed longer with Phosphor screens.</li>
</ol>

The authors have nothing to disclose.

Nanobodies represent an emerging class of protein binder scaffolds with many advantages over conventional antibodies: they are small, stable, monomeric, can be selected for high affinity and lack disulfide bonds33,35,44,45. They are used in a number of applications, such as in cell culture systems and organisms in developmental biology46,47</...

Retrograde traffic of proteins and lipids from the cell surface to various intracellular compartments is crucial for maintenance of membrane homeostasis to counterbalance secretion and to recycle components of anterograde transport machineries1,2. Following internalization via clathrin-dependent or -independent endocytosis, protein and lipid cargo first populate early endosomes from where they are further redirected either along the endo-lysosomal system, recycled to the plasma membrane, or targeted to the trans-Golgi network (TGN). Recycling from endosomes and/or the cell surface to the TGN is part of the functional cycle of a number of anterograde transmembrane cargo receptors, such as the cation-dependent and cation-independent mannose-6-phosphate receptors (CDMPR and CIMPR) delivering newly synthesized lysosomal hydrolases from the TGN to late endosomes and lysosomes3,4,5, sortilin and SorLA6,7, and Wntless (WLS) transporting Wnt ligands to the cell surface8,9,10,11. Other proteins retrieved back to the TGN are TGN46 and its related isoforms12,13,14, SNAREs (soluble N-ethylmaleimide-sensitive fusion factor attachment receptors)15,16,17, amyloid precursor protein (APP)18,19, progressive ankylosis (ANK) protein20, metal transporters such as ATP7A/B or DMT121,22, and transmembrane processing enzymes including carboxypeptidase D, furin or BACE123,24,25. Apart from these endogenous proteins, bacterial and plant toxins (e.g., Shiga and cholera toxin, ricin and abrin) hijack retrograde transport machineries to reach the ER for retrotranslocation into the cytosol26,27,28,29.
In order to directly analyze retrograde traffic, we have previously developed a nanobody-based toolkit to label and follow cargo proteins from the cell surface to intracellular compartments30. Nanobodies represent a new family of protein binders derived from homodimeric heavy-chain-only antibodies (hcAbs) that naturally occur in camelids and cartilaginous fishes31,32. They constitute the variable heavy-chain domain (VHH) of hcAbs and have many advantages over conventional antibodies (e.g., IgGs): They are monomeric, small (~15 kDa), highly soluble, devoid of disulfide bonds, can be bacterially expressed, and selected for high-affinity binding33,34,35,36. To make our nanobody tool versatile and broadly applicable, we employed functionalized anti-GFP nanobodies to surface-label and track proteins tagged with GFP at their extracellular/lumenal domain. By functionalization of nanobodies with mCherry, ascorbate peroxidase 2 (APEX2)37, or tyrosine sulfation (TS) sequences, retrograde transport of bonafide transmembrane cargo proteins can be analyzed by either fixed and live cell imaging, by electron microscopy, or biochemically. Since tyrosine sulfation mediated by tyrosylprotein sulfotransferases (TPST1 and TPST2) is a posttranslational modification restricted to the trans-Golgi/TGN, we can directly study transport and kinetics of proteins of interest from the cell surface to this intracellular Golgi compartment38,39,40.
In this methods article, we describe the ease of production of functionalized nanobodies (VHH-2xTS, -APEX2, -mCherry and derivatives) suited for a number of applications to analyze retrograde transport in mammalian cells30. We mainly focus on the use of TS site-modified nanobody for analysis of intracellular traffic from the cell surface to the compartment of sulfation.

<table >
	
		
		
		
		
	
	<tbody>
		<tr >
			<td >Anti-GFP antibody</td>
			<td >Sigma-Aldrich</td>
			<td >118144600001</td>
			<td >Product is distributed by Sigma-Aldrich, but manufactured by Roche</td>
		</tr>
		<tr >
			<td >Anti-His6 antibody</td>
			<td >Bethyl Laboratories</td>
			<td >A190-114A</td>
			<td ></td>
		</tr>
		<tr >
			<td >Anti-actin antibody</td>
			<td >EMD Millipore</td>
			<td >MAB1501</td>
			<td ></td>
		</tr>
		<tr >
			<td >Goat anti-rabbit HRP</td>
			<td >Sigma-Aldrich</td>
			<td >A-0545</td>
			<td ></td>
		</tr>
		<tr >
			<td >Goat anti-mouse HRP</td>
			<td >Sigma-Aldrich</td>
			<td >A-0168</td>
			<td ></td>
		</tr>
		<tr >
			<td >4&#39;,6-diamidino-2-phenylindole (DAPI)</td>
			<td >Sigma-Aldrich</td>
			<td >D9542</td>
			<td >dissolved in 1 x PBS/1%BSA</td>
		</tr>
		<tr >
			<td >Dimethyl sulfoxide (DMSO)</td>
			<td >Applichem</td>
			<td >A3672</td>
			<td ></td>
		</tr>
		<tr >
			<td >D-biotin</td>
			<td >Sigma-Aldrich</td>
			<td >B4501</td>
			<td >dissolved in sterile 500 mM NaH2PO4 or DMSO</td>
		</tr>
		<tr >
			<td >5-aminolevuilnic acid (dALA) hydrochloride</td>
			<td >Sigma-Aldrich</td>
			<td >A3785</td>
			<td >dissolved in sterile water</td>
		</tr>
		<tr >
			<td >DNase I</td>
			<td >Applichem</td>
			<td >A3778</td>
			<td >dissolved in sterile water</td>
		</tr>
		<tr >
			<td >Lysozyme</td>
			<td >Sigma-Aldrich</td>
			<td >18037059001</td>
			<td >Product is distributed by Sigma-Aldrich, but manufactured by Roche</td>
		</tr>
		<tr >
			<td >Brefeldin A (BFA)</td>
			<td >Sigma-Aldrich</td>
			<td >B5936</td>
			<td ></td>
		</tr>
		<tr >
			<td >Puromycin</td>
			<td >Invivogen</td>
			<td >ant-pr-1</td>
			<td ></td>
		</tr>
		<tr >
			<td >Penicillin/Streptomycin</td>
			<td >Bioconcept</td>
			<td >4-01F00-H</td>
			<td ></td>
		</tr>
		<tr >
			<td >L-glutamine</td>
			<td >Applichem</td>
			<td >A3704</td>
			<td ></td>
		</tr>
		<tr >
			<td >Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM)</td>
			<td >Sigma-Aldrich</td>
			<td >D5796</td>
			<td ></td>
		</tr>
		<tr >
			<td >Fetal calf serum (FCS)</td>
			<td >Biowest</td>
			<td >S181B-500</td>
			<td ></td>
		</tr>
		<tr >
			<td >Sulfur-35 as sodium sulfate</td>
			<td >Hartmann Analytics</td>
			<td >ARS0105</td>
			<td >Product contains 5 mCi</td>
		</tr>
		<tr >
			<td >Earle&#39;s balanced salts</td>
			<td >Sigma-Aldrich</td>
			<td >E6267</td>
			<td ></td>
		</tr>
		<tr >
			<td >MEM amino acids (50 x) solution</td>
			<td >Sigma-Aldrich</td>
			<td >M5550</td>
			<td ></td>
		</tr>
		<tr >
			<td >MEM vitamin solution (100 x)</td>
			<td >Sigma-Aldrich</td>
			<td >M6895</td>
			<td ></td>
		</tr>
		<tr >
			<td >cOmplete, Mini Protease inhibitor cocktail</td>
			<td >Sigma-Aldrich</td>
			<td >11836153001</td>
			<td >Product is distributed by Sigma-Aldrich, but manufactured by Roche</td>
		</tr>
		<tr >
			<td >Isopropyl-&#946;-D-thiogalactopyranosid (IPTG)</td>
			<td >Applichem</td>
			<td >A1008</td>
			<td >dissolved in sterile water, stock is 1 M</td>
		</tr>
		<tr >
			<td >Carbenicillin disodium salt</td>
			<td >Applichem</td>
			<td >A1491</td>
			<td >dissolved in sterile water, stock is 100 mg/mL</td>
		</tr>
		<tr >
			<td >Kanamycin sulfate</td>
			<td >Applichem</td>
			<td >A1493</td>
			<td >dissolved in sterile water, stock is 100 mg/mL</td>
		</tr>
		<tr >
			<td >Coomassie-R (Brilliant Blue)</td>
			<td >Sigma-Aldrich</td>
			<td >B-0149</td>
			<td ></td>
		</tr>
		<tr >
			<td >Paraformaldehyde (PFA)</td>
			<td >Applichem</td>
			<td >A3813</td>
			<td ></td>
		</tr>
		<tr >
			<td >Bovine serum albumin (BSA)</td>
			<td >Sigma-Aldrich</td>
			<td >A2153</td>
			<td ></td>
		</tr>
		<tr >
			<td >Fluoromount-G</td>
			<td >Southern Biotech</td>
			<td >0100-01</td>
			<td ></td>
		</tr>
		<tr >
			<td >Ni Sepharose High Performance</td>
			<td >GE Healthcare</td>
			<td >17-5268-01</td>
			<td ></td>
		</tr>
		<tr >
			<td >His GraviTrap columns</td>
			<td >GE Healthcare</td>
			<td >GE11-0033-99</td>
			<td ></td>
		</tr>
		<tr >
			<td >His buffer kit</td>
			<td >GE Healthcare</td>
			<td >GE11-0034-00</td>
			<td ></td>
		</tr>
		<tr >
			<td >Disposable PD10 desalting columns</td>
			<td >GE Healthcare</td>
			<td >GE17-0851-01</td>
			<td ></td>
		</tr>
		<tr >
			<td >Mini-Protean TGX gels, 4-20%, 15-well</td>
			<td >Bio-Rad</td>
			<td >456-1096</td>
			<td ></td>
		</tr>
		<tr >
			<td >Dulbecco&#8217;s phosphate buffered saline (DPBS) w/o Ca2+/Mg2+</td>
			<td >Sigma-Aldrich</td>
			<td >D8537</td>
			<td ></td>
		</tr>
		<tr >
			<td >35-mm dishes</td>
			<td >Falcon</td>
			<td >353001</td>
			<td ></td>
		</tr>
		<tr >
			<td >6-well plates</td>
			<td >TPP</td>
			<td >92406</td>
			<td ></td>
		</tr>
		<tr >
			<td >Glass coverslips (No. 1.5H)</td>
			<td >VWR</td>
			<td >631-0153</td>
			<td ></td>
		</tr>
		<tr >
			<td >Phenylmethylsulfonyl fluoride (PMSF)</td>
			<td >Applichem</td>
			<td >A0999.0025</td>
			<td >dissolved in 40% DMSO 60% isopropanol, stock in 500 mM</td>
		</tr>
		<tr >
			<td >Tryptone</td>
			<td >Applichem</td>
			<td >A1553</td>
			<td ></td>
		</tr>
		<tr >
			<td >Yeast extract</td>
			<td >Applichem</td>
			<td >A1552</td>
			<td></td>
		</tr>
		<tr >
			<td >Magnesium chloride hexahydrate</td>
			<td >Merck Millipore</td>
			<td >105833</td>
			<td>dissolved in sterile water, stock is 1 M</td>
		</tr>
		<tr >
			<td >Calcium chloride dihydrate</td>
			<td >Merck Millipore</td>
			<td >102382</td>
			<td>dissolved in sterile water, stock is 1 M</td>
		</tr>
		<tr >
			<td >Sodium chloride</td>
			<td >Merck Millipore</td>
			<td >106404</td>
			<td>dissolved in sterile water, stock is 5 M</td>
		</tr>
	</tbody>
</table>

analysis of endocytic uptake and retrograde transport to the trans-golgi network using functionalized nanobodies in cultured cells

Transport of proteins and membranes from the cell surface to the Golgi and beyond is essential for homeostasis, organelle identity and physiology. To study retrograde protein traffic, we have recently developed a versatile nanobody-based toolkit to analyze transport from the cell surface to the Golgi complex, either by fixed and live cell imaging, by electron microscopy, or biochemically. We engineered functionalized anti-green fluorescent protein (GFP) nanobodies &#8212; small, monomeric, high-affinity protein binders &#8212; that can be applied to cell lines expressing membrane proteins of interest with an extracellular GFP moiety. Derivatized nanobodies bound to the GFP reporters are specifically internalized and transported piggyback along the reporters&#39; sorting routes. Nanobodies were functionalized with fluorophores to follow retrograde transport by fluorescence microscopy and live imaging, with ascorbate peroxidase 2 (APEX2) to investigate the ultrastructural localization of reporter-nanobody complexes by electron microscopy, and with tyrosine sulfation (TS) motifs to assess kinetics of trans-Golgi network (TGN) arrival. In this methodological article, we outline the general procedure to bacterially express and purify functionalized nanobodies. We illustrate the powerful use of our tool using the mCherry- and TS-modified nanobodies to analyze endocytic uptake and TGN arrival of cargo proteins.

To investigate retrograde protein transport to various intracellular destinations, we have recently established an anti-GFP nanobody-based tool to label and follow recombinant fusion proteins from the cell surface30. Here, we demonstrate the bacterial production of such derivatized nanobodies and demonstrate their application to study endocytic uptake by fluorescence microscopy and immunoblotting, as well as their use to investigate TGN arrival by sulfation analysi...

Watch this Scientific Journal Video about Analysis of Endocytic Uptake and Retrograde Transport to the Trans-Golgi Network Using Functionalized Nanobodies in Cultured Cells at JoVE.com

Analysis of Endocytic Uptake and Retrograde Transport to the Trans-Golgi Network Using Functionalized Nanobodies in Cultured Cells

Retrograde transport of proteins from the cell surface to the Golgi is essential to maintain membrane homeostasis. Here, we describe a method to biochemically analyze cell surface-to-Golgi transport of recombinant proteins using functionalized nanobodies in HeLa cells.

Transport of proteins and membranes from the cell surface to the Golgi and beyond is essential for homeostasis, organelle identity and physiology. To ...

This method enables the production and application of bacterial expressed, functionalised nano-bodies to analyze retrograde protein transport from the cell surface to intercellular compartments.In particular the Trans-Golgi network.The main advantage of this technique is the use of sulfation.A highly specific, compartmentalized, and sensitive post-translation modification to assess reporter nano-body complex arrival in the DTN compartment.This method in combination with other derivatised nano-bodies for life cell imaging and electron microscopy can also be used to study endocitic processes of disease-related cell surface receptors.Our repertoire of functionalised nano-bodies has been engineered to study retrograde transport in cultured cells.However, these tools may also be applied in multicellular organisms.To begin this procedure, pick a bacterial colony containing the vector of interest from the plate and inoculate a flask containing 20 milliliters of LB medium supplemented with antibiotics.Incubate overnight at 37

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Intracellular Membrane Traffic

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8.9K vistas.  University of Basel. Este método permite la producción y aplicación de nanosí cuerpos funcionalizados y expresados bacterianos para analizar el transporte de proteínas retrógradas desde la superficie celular hasta compartimentos intercelulares. En particular, la red Trans-Golgi. La principal ventaja de esta técnica es el uso de la sulfatación. Una modificación altamente específica, compartimentada y sensible después de la traducción para evaluar la llegada del complejo de nano-cuerpo de reportero en e...

Video: Analysis of Endocytic Uptake and Retrograde Transport to the Trans-Golgi Network Using Functionalized Nanobodies in Cultured Cells

Nitrogen Cavitation and Differential Centrifugation Allows for Monitoring the Distribution of Peripheral Membrane Proteins in Cultured Cells

Cultured cells are useful for studying the subcellular distribution of proteins, including peripheral membrane proteins. Genetically encoded fluorescently tagged proteins have revolutionized the study of subcellular protein distribution. However, it is difficult to quantify the distribution with fluorescent microscopy, especially when proteins are partially cytosolic. Moreover, it is often important to study endogenous proteins. Biochemical assays such as immunoblots remain the gold standard for quantification of protein distribution after subcellular fractionation. Although there are commercial kits that aim to isolate cytosolic or certain membrane fractions, most of these kits are based on extraction with detergents, which may be unsuitable for studying peripheral membrane proteins that are easily extracted from membranes. Here we present a detergent-free protocol for cellular homogenization by nitrogen cavitation and subsequent separation of cytosolic and membrane-bound proteins by ultracentrifugation. We confirm the separation of subcellular organelles in soluble and pellet fractions across different cell types, and compare protein extraction among several common non-detergent-based mechanical homogenization methods. Among several advantages of nitrogen cavitation is the superior efficiency of cellular disruption with minimal physical and chemical damage to delicate organelles. Combined with ultracentrifugation, nitrogen cavitation is an excellent method to examine the shift of peripheral membrane proteins between cytosolic and membrane fractions.

Here we present protocols for detergent-free homogenization of cultured mammalian cells based on nitrogen cavitation and subsequent separation of cytosolic and membrane-bound proteins by ultracentrifugation. This method is ideal for monitoring the partitioning of peripheral membrane proteins between soluble and membrane fractions.

Cultured cells are useful for studying the subcellular distribution of proteins, including peripheral membrane proteins. Genetically encoded fluorescently ...

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

Proofreading and DNA Repair Assay Using Single Nucleotide Extension and MALDI-TOF Mass Spectrometry Analysis

The maintenance of the genome and its faithful replication is paramount for conserving genetic information. To assess high fidelity replication, we have developed a simple non-labeled and non-radio-isotopic method using a matrix-assisted laser desorption ionization with time-of-flight (MALDI-TOF) mass spectrometry (MS) analysis for a proofreading study. Here, a DNA polymerase [e.g., the Klenow fragment (KF) of Escherichia coli DNA polymerase I (pol I) in this study] in the presence of all four dideoxyribonucleotide triphosphates is used to process a mismatched primer-template duplex. The mismatched primer is then proofread/extended and subjected to MALDI-TOF MS. The products are distinguished by the mass change of the primer down to single nucleotide variations. Importantly, a proofreading can also be determined for internal single mismatches, albeit at different efficiencies. Mismatches located at 2-4-nucleotides (nt) from the 3' end were efficiently proofread by pol I, and a mismatch at 5 nt from the primer terminus showed only a partial correction. No proofreading occurred for internal mismatches located at 6 - 9 nt from the primer 3' end. This method can also be applied to DNA repair assays (e.g., assessing a base-lesion repair of substrates for the endo V repair pathway). Primers containing 3' penultimate deoxyinosine (dI) lesions could be corrected by pol I. Indeed, penultimate T-I, G-I, and A-I substrates had their last 2 dI-containing nucleotides excised by pol I before adding a correct ddN 5'-monophosphate (ddNMP) while penultimate C-I mismatches were tolerated by pol I, allowing the primer to be extended without repair, demonstrating the sensitivity and resolution of the MS assay to measure DNA repair.

A non-labeled, non-radio-isotopic method for DNA polymerase proofreading and a DNA repair assay was developed by using high-resolution MALDI-TOF mass spectrometry and a single nucleotide extension strategy. The assay proved to be very specific, simple, rapid, and easy to perform for proofreading and repair patches shorter than 9-nucleotides.

The maintenance of the genome and its faithful replication is paramount for conserving genetic information. To assess high fidelity replication, we have ...

Dissipative Microgravimetry to Study the Binding Dynamics of the Phospholipid Binding Protein Annexin A2 to Solid-supported Lipid Bilayers Using a Quartz Resonator

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic properties of the absorbed/immobilized mass on sensor surfaces, allowing the measurements of the interaction of proteins with solid-supported surfaces, such as lipid bilayers, in real-time and with a high sensitivity. Annexins are a highly conserved group of phospholipid-binding proteins that interact reversibly with the negatively charged headgroups via the coordination of calcium ions. Here, we describe a protocol that was employed to quantitatively analyze the binding of annexin A2 (AnxA2) to planar lipid bilayers prepared on the surface of a quartz sensor. This protocol is optimized to obtain robust and reproducible data and includes a detailed step-by-step description. The method can be applied to other membrane-binding proteins and bilayer compositions.

Here, we present an experimental protocol that can be employed to determine the binding affinities and mode of interaction of label-free phospholipid-binding protein annexin A2 with immobilized solid-supported bilayers (SLB) by simultaneously measuring the mass uptake and the viscoelastic properties of the protein annexin A2.

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic ...

Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and the degradation of target proteins in live cells. By using short (pulse) radiolabeling times (&lt;30 min) and tightly controlled chase times, it is possible to label only a small fraction of the total protein pool and follow its folding. When combined with nonreducing/reducing SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoprecipitation with (conformation-specific) antibodies, folding processes can be examined in great detail. This system has been used to analyze the folding of proteins with a huge variation in properties such as soluble proteins, single and multi-pass transmembrane proteins, heavily N- and O-glycosylated proteins, and proteins with and without extensive disulfide bonding. Pulse-chase methods are the basis of kinetic studies into a range of additional features, including co- and posttranslational modifications, oligomerization, and polymerization, essentially allowing the analysis of a protein from birth to death. Pulse-chase studies on protein folding are complementary with other biochemical and biophysical methods for studying proteins in vitro by providing increased temporal resolution and physiological information. The methods as described within this paper are adapted easily to study the folding of almost any protein that can be expressed in mammalian or insect-cell systems.

Here we describe a protocol for a general pulse-chase method that allows the kinetic analysis of folding, transport, and degradation of proteins to be followed in live cells.

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and ...

A Strategy to Identify Compounds that Affect Cell Growth and Survival in Cultured Mammalian Cells at Low-to-Moderate Throughput

Cytotoxicity is a critical parameter that needs to be quantified when studying drugs that may have therapeutic benefits. Because of this, many drug screening assays utilize cytotoxicity as one of the critical characteristics to be profiled for individual compounds. Cells in culture are a useful model to assess cytotoxicity before proceeding to follow up on promising lead compounds in more costly and labor-intensive animal models. We describe a strategy to identify compounds that affect cell growth in a tdTomato expressing human neural stem cells (NSC) line. The strategy uses two complementary assays to assess cell number. One assay works via the reduction of 3-(4,5-dimethylthizol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) to formazan as a proxy for cell number and the other directly counts the tdTomato expressing NSCs. The two assays can be performed simultaneously in a single experiment and are not labor intensive, rapid, and inexpensive. The strategy described in this demonstration tested 57 compounds in an exploratory primary screen for toxicity in a 96-well plate format. Three of the hits were characterized further in a six-point dose response using the same assay set-up as the primary screen. In addition to providing excellent corroboration for toxicity, comparison of results from the two assays may be effective in identifying compounds affecting other aspects of cell growth.

It is often necessary to assess the potential cytotoxicity of a set of compounds on cultured cells. Here, we describe a strategy to reliably screen for toxic compounds in a 96-well format.

Cytotoxicity is a critical parameter that needs to be quantified when studying drugs that may have therapeutic benefits. Because of this, many drug ...

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

Density Gradient Ultracentrifugation for Investigating Endocytic Recycling in Mammalian Cells

Endosomal trafficking is an essential cellular process that regulates a broad range of biological events. Proteins are internalized from the plasma membrane and then transported to the early endosomes. The internalized proteins could be transited to the lysosome for degradation or recycled back to the plasma membrane. A robust endocytic recycling pathway is required to balance the removal of membrane materials from endocytosis. Various proteins are reported to regulate the pathway, including ADP-ribosylation factor 6 (ARF6). Density gradient ultracentrifugation is a classical method for cell fractionation. After the centrifugation, organelles are sedimented at their isopycnic surface. The fractions are collected and used for other downstream applications. Described here is a protocol to obtain a recycling endosome-containing fraction from transfected mammalian cells using density gradient ultracentrifugation. The isolated fractions were subjected to standard Western blotting for analyzing their protein contents. By employing this method, we identified that the plasma membrane targeting of engulfment and cell motility 1 (ELMO1), a Ras-related C3 botulinum toxin substrate 1 (Rac1) guanine nucleotide exchange factor, is through ARF6-mediated endocytic recycling.

This paper aims to present a protocol for preparing recycling endosomes from mammalian cells using sucrose density gradient ultracentrifugation.

Endosomal trafficking is an essential cellular process that regulates a broad range of biological events. Proteins are internalized from the plasma ...

JUMPn: A Streamlined Application for Protein Co-Expression Clustering and Network Analysis in Proteomics

With recent advances in mass spectrometry-based proteomics technologies, deep profiling of hundreds of proteomes has become increasingly feasible. However, deriving biological insights from such valuable datasets is challenging. Here we introduce a systems biology-based software JUMPn, and its associated protocol to organize the proteome into protein co-expression clusters across samples and protein-protein interaction (PPI) networks connected by modules (e.g., protein complexes). Using the R/Shiny platform, the JUMPn software streamlines the analysis of co-expression clustering, pathway enrichment, and PPI module detection, with integrated data visualization and a user-friendly interface. The main steps of the protocol include installation of the JUMPn software, the definition of differentially expressed proteins or the (dys)regulated proteome, determination of meaningful co-expression clusters and PPI modules, and result visualization. While the protocol is demonstrated using an isobaric labeling-based proteome profile, JUMPn is generally applicable to a wide range of quantitative datasets (e.g., label-free proteomics). The JUMPn software and protocol thus provide a powerful tool to facilitate biological interpretation in quantitative proteomics.

We present a systems biology tool JUMPn to perform and visualize network analysis for quantitative proteomics data, with a detailed protocol including data pre-processing, co-expression clustering, pathway enrichment, and protein-protein interaction network analysis.

With recent advances in mass spectrometry-based proteomics technologies, deep profiling of hundreds of proteomes has become increasingly feasible. ...

Routine Collection of High-Resolution cryo-EM Datasets Using 200 KV Transmission Electron Microscope

Cryo-electron microscopy (cryo-EM) has been established as a routine method for protein structure determination during the past decade, taking an ever-increasing share of published structural data. Recent advances in TEM technology and automation have boosted both the speed of data collection and quality of acquired images while simultaneously decreasing the required level of expertise for obtaining cryo-EM maps at sub-3 &#197; resolutions. While most of such high-resolution structures have been obtained using state-of-the-art 300 kV cryo-TEM systems, high-resolution structures can be also obtained with 200 kV cryo-TEM systems, especially when equipped with an energy filter. Additionally, automation of microscope alignments and data collection with real-time image quality assessment reduces system complexity and assures optimal microscope settings, resulting in increased yield of high-quality images and overall throughput of data collection. This protocol demonstrates the implementation of recent technological advances and automation features on a 200 kV cryo-transmission electron microscope and shows how to collect data for the reconstruction of 3D maps that are sufficient for de novo atomic model building. We focus on best practices, critical variables, and common issues that must be considered to enable the routine collection of such high-resolution cryo-EM datasets. Particularly the following essential topics are reviewed in detail: i) automation of microscope alignments, ii) selection of suitable areas for data acquisition, iii) optimal optical parameters for high-quality, high-throughput data collection, iv) energy filter tuning for zero-loss imaging, and v) data management and quality assessment. Application of the best practices and improvement of achievable resolution using an energy filter will be demonstrated on the example of apo-ferritin that was reconstructed to 1.6 &#197;, and Thermoplasma acidophilum 20S proteasome reconstructed to 2.1-&#197; resolution using a 200 kV TEM equipped with an energy filter and a direct electron detector.

High-resolution cryo-EM maps of macromolecules can be also achieved by using 200 kV TEM microscopes. This protocol shows the best practices for setting accurate optics alignments, data acquisition schemes, and selection of imaging areas that are all essential for the successful collection of high-resolution datasets using a 200 kV TEM.