Raz Zarivach is supported by the Israel Science Foundation (grant no. 163/22). Tomer Eli Ben Yosef and Arie Moran are supported by the Israel Science Foundation (grant no. 2047/20). We would like to thank Daniel Gitler and his group at Ben-Gurion University for their cooperation, support, and expertise.

1. Cell transfection
<ol>
	<li>Culture HEK293T cells in Dulbecco&#39;s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, and 1x penicillin/streptomycin (see Table of Materials) in a humidified incubator at 37 &#176;C/5% CO2 until confluence on a 10 cm plate (8.8 x 106 total cells).</li>
	<li>Place one 13 mm coverslip in each of the wells of a 12-well plate. Dilute 0.44 x 106 trypsinized cells from step 1.1 in 12 mL of complete DMEM. Mix well by pipetting up and down three to five times. Fill each well with 1 mL of the mixed solution. Grow in a humidified incubator at 37 &#176;C/5% CO2 overnight.</li>
	<li>Replace the complete DMEM in each well with 1 mL of serum-free DMEM (see Table of Materials) in each well. Return the 12-well plate to the humidified incubator as in step 1.1.</li>
	<li>For each transfection well, dilute 1 &#181;g of pAAV2 plasmid containing the protein of interest tagged with mCherry fluorescent protein (Supplementary File 1) in 100 &#181;L of serum-free DMEM in a 1.5 mL tube.</li>
	<li>Add 3 &#181;g of polyethyleneimine (PEI, see Table of Materials) to 100 &#181;L of serum-free DMEM per transfection, with each transfection in a different 1.5 mL tube. The plasmid:PEI ratio should be 1:3 (&#181;g:&#181;g).</li>
	<li>Vortex both tubes from step 1.4&#160;and step 1.5&#160;for 10 s, and leave to rest at room temperature for at least 5 min.</li>
	<li>Mix one part (100 &#181;L per well) of the DNA solution from step 1.4 with one part (100 &#181;L per well) of the PEI solution from step 1.5. Vortex the final solution for 10 s, and leave it to rest at room temperature for at least 20 min and for up to 6 h.</li>
	<li>Take the 12-well plate from the incubator. Vortex the final solution from step 1.7, and add 200 &#181;L to each of the wells in the 12-well plate from step 1.3. Return the 12-well plate to the humidified incubator (step 1.1).</li>
	<li>After 3 h, replace the medium in each well with 1 mL of complete DMEM. Return the 12-well plate to the humidified incubator as in step 1.1.</li>
	<li>After 2 days, take the transfected cell culture from the 37 &#176;C/5% CO2 incubator, and place it on an inverted fluorescence microscope using 10x magnification.</li>
	<li>Using the microscope focus wheel, focus on the cells while using brightfield light.</li>
	<li>Switch to the fluorescent mCherry excitation (587 nm) and emission (610 nm) wavelengths, and turn off the brightfield light. Check for the fluorescence of the cells to confirm the expression of ZnT1 mCherry.</li>
	<li>Prepare the dye loading solution.
	<ol>
		<li>Take an aliquot of 4 &#181;L of zinc-specific fluorescent dye (see Table of Materials) in its acetoxymethyl (AM) ester form, dissolved in DMSO (1 &#181;g/&#181;L), from a light-protected stock stored in a &#8722;20&#160;&#176;C freezer. This form allows the dye to enter the cell through the plasma membrane by simple diffusion.</li>
		<li>Add 4 &#181;L of 10% pluronic acid (or 2 &#181;L of 20% pluronic acid, see Table of Materials) to the aliquot from step 1.13.1. Mix the solution well by pipetting up and down three times, and then add all 8 &#181;L to a 1.5 mL tube.</li>
		<li>Add 750 &#181;L of Ringer&#39;s solution (prepared in-house, see Table of Materials for composition) supplemented with 1 mg/mL (~0.1%) bovine serum albumin to the tube from step 1.13.2, and vortex vigorously. Add another 750 &#181;L of the same solution, and vortex again to ensure maximum mixing.</li>
	</ol>
	</li>
	<li>Add 750 &#181;L of the final solution from step 1.13.3&#160;to two wells in a new 6-well culture plate. Cover with aluminum foil. 
	NOTE: To avoid bleaching, the 6-well plate is always covered with aluminum foil from this step.</li>
	<li>Using a fine tweezer, take up to four replicate coverslips from the transfected cell culture plate, and place them in the first filled well (up to four slides per well) of the new 6-well plate from step 1.14. Repeat this process for the second filled well. 
	NOTE: All the coverslips in the same filled well must be from the same condition. However, each well can contain different conditions.</li>
	<li>Cover with aluminum foil, and gently shake for 15-20 min.</li>
	<li>Remove the dye loading solution, and replace it with a new washing solution of Ringer&#39;s plus albumin, as used in&#160;step 1.13.3. Cover with aluminum foil. Leave to shake again for 20 min. This allows the cleavage of the AM ester by the intracellular esterases.</li>
</ol>
2. Microscope preparation
<ol>
	<li>Arrange the necessary tools (see Table of Materials) as mentioned: an inverted fluorescence microscope capable of detecting GFP and mCherry fluorophores, a perfusion system that allows switching between at least two solutions, a suction system, and a perfusion chamber.</li>
	<li>Turn on the microscope, its light source, and the camera. Turn on the suction system.</li>
	<li>Wash the perfusion system. Wash the first chamber with Ringer&#39;s solution and the second chamber with Ringer&#39;s solution containing 7 &#181;M zinc solution supplemented with 7 &#181;M pyrithione (zinc ionophore, see Table of Materials). To avoid air bubbles or gaps, leave some liquid in each chamber. 
	NOTE: Since both containers are connected to the same tube hooked to the perfusion chamber, always ensure that the shared segment is washed with Ringer&#39;s solution.</li>
	<li>Close the taps, and fill the appropriate chambers with Ringer&#39;s solution and Ringer&#39;s zinc solution, as in step 2.3.</li>
</ol>
3. Sample preparation
<ol>
	<li>Take the perfusion chamber, and place it with the narrow side of the groove facing upward. 
	NOTE: The groove is a hole in the middle of the perfusion chamber. It allows the perfusion fluid access to the cells. One side of the groove is narrow, and the opposite side is wide.</li>
	<li>Apply a sealing silicone (see Table of Materials) around the groove. Clean any sealing silicone from the groove using a pipette tip. 
	NOTE: Ensure there is enough silicone seal around the groove to seal a 22 mm coverslip.</li>
	<li>Using a fine tweezer, take a coverslip from the washing solution, and place it on top of the groove with the cells facing down. This way, the cells will be exposed to the solution perfusing the groove during the experiment.</li>
	<li>Place a 22 mm coverslip on top of the 13 mm coverslip, and tighten it using the tweezer. Take care not to crack the coverslips.</li>
	<li>Flip the chamber, and press on it to release all the present liquids. Fill the groove with 100 &#181;L of Ringer&#39;s washing solution, and press again to ensure no leakage. 
	NOTE: If a leakage is detected, apply a sealing agent at the site of the leakage, and retest.</li>
	<li>Mount the perfusion chamber onto the platform, and secure it. Place the perfusion and suction tubes to allow perfusion over the cells in the groove.</li>
	<li>Change the perfusion rate to approximately 2 mL/min, and turn on the perfusion of the Ringer&#39;s solution. Ensure the perfusion system is working with no leakage or spillover.</li>
</ol>
4. Measurement preparation
<ol>
	<li>Open the imaging software (see Table of Materials) by double-clicking on the icon. Log in with the relevant credentials. Choose the attached camera, and press Ok.</li>
	<li>Set the microscope magnification to 10x using the button on the left side of the microscope.</li>
	<li>Choosing live view and using the joystick, move the platform to focus on the cells in the groove.</li>
	<li>While the perfusion is on, turn off the lights and change the wavelength to mCherry by pressing the dedicated button in the interface. Adjust the focus using the microscope focus wheel.</li>
	<li>Once the microscope is focused on the cells, move the platform using the joystick to allow the selection of the appropriate cell patches. 
	NOTE: A suitable area is considered an area with at least 10 cells for an ROI and an empty area for background.</li>
	<li>Select the Turn&#160;ROIs On/Off drop-down menu, and choose Draw Circular ROIs.</li>
	<li>Draw ROIs of cell clusters with the mCherry-expressing cells (indicates the expression of ZnT1).</li>
	<li>From the same toolbar as step 4.6, click the Turn background ROIs On/Off button, and adjust the location and size of the background ROI to an area with no cells at all.
	<ol>
		<li>Check with the mCherry and EGFP wavelengths to ensure no cells are in the selected background ROI. 
		NOTE: The wavelengths were adjusted so that mCherry used an excitation wavelength of 520 nm and an emission wavelength of 610 nm and EGFP used an excitation wavelength of 470 nm and an emission wavelength of 520 nm.</li>
	</ol>
	</li>
	<li>Select the Wavelength (&#955;) sub-tab. Mark its checkbox, and ensure that the GFP wavelength is present and its checkbox marked. If not, add and mark it.</li>
	<li>Select the Duration sub-tab. Define the measurement interval as every 5 s, and change the number of intervals to suit the experiment duration.</li>
	<li>In the Focus section on the microscope panel below the eyepiece, click on the On button to enable the perfect focus system (PFS).</li>
	<li>Using the PFS focus wheel, adjust the focus. In the software interface, Under ND Acquisition, ensure PFS on is ticked.</li>
	<li>Click on Run now.</li>
</ol>
5. Experimental procedure
<ol>
	<li>Start with a 90 s baseline period measurement using Ringer&#39;s solution perfusion.</li>
	<li>After the baseline period ends, turn off the Ringer&#39;s solution perfusion, and then turn on the Ringer&#39;s zinc solution perfusion. Mark the switching of the solution by clicking the red flag on the bottom right of the main interface panel. A rise in fluorescence is expected to appear.</li>
	<li>Once the fluorescence rise starts to saturate, change back to perfusion with Ringer&#39;s solution. Turn off the Ringer&#39;s zinc solution perfusion, and then turn on the Ringer&#39;s solution perfusion.</li>
	<li>Wait until the experiment time expires. A noticeable yet steady decrease in fluorescence is expected if the ZnT1 is working properly.</li>
</ol>
6. Data export
<ol>
	<li>To export the data with baseline subtraction, press the Subtract Baseline button, and view the changes in the &#34;ND acquisition window&#34;.</li>
	<li>Click on the Export button in the software interface in the &#34;ND acquisition window&#34;. An excel datasheet will open. Save to the desired location.</li>
</ol>
7. Data analysis
<ol>
	<li>For each ROI, create an average baseline fluorescence from the first 90-100 s.</li>
	<li>Express the fluorescence calculated for each ROI as a percentage of the background for that ROI.</li>
	<li>Create a row average of all the ROIs, and plot the result as a line graph. This creates an average of all the ROIs as a function of time.</li>
	<li>Using the linear fit function, select the initial rate for the decrease in fluorescence following the wash with Ringer&#39;s solution. The slope value of the equation correlates with the transport rate.</li>
</ol>

Investigating Zinc Transport Mechanism in Mammalian Cells with FluoZin-3 Assay

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The authors declare no conflicts of interest.

The above-described method allows for the direct measurement of the intracellular zinc concentration with high temporal resolution. Compared to other methods, this method involving monitoring changes in intracellular Zn2+ can substantially decrease background noise. In addition, the dye&#39;s selectivity for zinc eliminates potential cross-interactions with other metal cations18,19. Finally, its lack of immediate cytotoxicity enables the testing of liv...

Zinc is an essential trace element in the cellular milieu. It incorporates one-third of all proteins and is involved in various cellular processes, such as catalysis1, transcription2, and structural motifs3. However, despite being redox-inert, high zinc concentrations are toxic to the cell, which is why no mammalian organism has survived without the presence of mechanisms regulating zinc homeostasis. In mammals, three mechanisms are responsible for this process: (1) metallothioneins, which are cytosolic cysteine-rich proteins that bind zinc at a high affinity, thus preventing excess free cytosolic zinc4; (2) Zrt/Irt-like proteins (ZIPs), which are zinc transporters responsible for zinc influx into the cytosol through the plasma membrane or from intracellular organelles4,5,6,7,8; and (3) ZnTs, which are a mammalian subset of the ubiquitous cation diffusion facilitator (CDF) family and are zinc transporters, as they extrude zinc from the cytosol across the plasma membrane or into the intracellular organelles4,5,6,7,8,9. Due to the importance of zinc to cellular metabolism, it is vital to understand cellular zinc dynamics.
Previous methods to assess zinc dynamics depended on assessing the expression levels of mRNA under different zinc conditions by correlating them with cellular zinc measurements of fixed tissues or cells10,11,12. These methods include chemical detection and immunohistochemistry staining. However, these methods yield only indirect measures and, thus, determine only an offline correlation between intracellular zinc concentration and the expression of zinc transporters. Consequently, these methods cannot infer any parameters requiring high temporal resolution.
A more direct measurement of Zn2+ transport uses radioactive isotopes of zinc13. This method relies on the measurement of radiolabeled Zn2+ to monitor zinc transport and its kinetics. However, due to the importance of zinc to cellular homeostasis, multiple cellular processes regulate intracellular zinc concentration. Among these are extracellular binding and several transport systems that work in concert to maintain tight control of intracellular Zn2+ levels. The combination of these processes creates considerable background noise, which makes it difficult to test individual zinc-related transport functions.
This article demonstrates a method to directly monitor the zinc transport rate by measuring the intracellular free zinc concentration using a zinc-specific fluorescent dye, FluoZin-3. The dye has high specificity for Zn2+ and little interference from other divalent cations, such as calcium. In addition, in its ester form, it enters the cells by nonionic diffusion and is then trapped due to the activity of intracellular di-esterase. Thus, its fluorescence is correlated primarily with the free cytosolic zinc concentration. These experiments were conducted to study the structure-function relationship of zinc transporter 1 (ZnT1), a member of the ZnT family.

<table><tbody><tr><td>10 cm plate</td><td>greiner bio-one</td><td>664160</td><td></td></tr><tr><td>12-well cell culture plate</td><td>greiner bio-one</td><td>665180</td><td></td></tr><tr><td>13 mm coverslips</td><td>Superior Marienfeld</td><td>111530</td><td></td></tr><tr><td>22 mm cover slides</td><td>Superior Marienfeld</td><td>101050</td><td></td></tr><tr><td>6-well culture plate</td><td>greiner bio-one</td><td>657160</td><td></td></tr><tr><td>Bovine serum albumin</td><td>bioWorld</td><td>22070008</td><td></td></tr><tr><td>Calcium chloride anhydrous, granular</td><td>Sigma Aldrich</td><td>C1016</td><td>Concentration in Ringer solution: 1 mM</td></tr><tr><td>D-(+)-Glucose</td><td>Glentham Life Science</td><td>GC6947</td><td>Concentration in Ringer solution: 10 mM</td></tr><tr><td>Dubelco&amp;rsquo;s Modified Eagle Media (DMEM)&amp;nbsp;</td><td>Sartorius</td><td>01-055-1A</td><td></td></tr><tr><td>Eclipse Ti inverted microscope</td><td>Nikon</td><td>TI-DH</td><td>Discontinued. Replaced by Eclipse Ti2</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Cytiva</td><td>SH30088.03</td><td></td></tr><tr><td>Fine tweezers</td><td>Dumont</td><td>0203-55-PS</td><td></td></tr><tr><td>Fluozin-3AM</td><td>Invitrogen</td><td>F24195</td><td></td></tr><tr><td>HyClone Penicillin-Streptomycin 100x solution</td><td>Cytiva</td><td>SV30010&amp;nbsp;</td><td></td></tr><tr><td>LED illumination system</td><td>CoolLED</td><td>pE-4000</td><td></td></tr><tr><td>L-glutamine</td><td>Biological Industries</td><td>03-020-1B</td><td></td></tr><tr><td>Magnesium chloride hexahydrate</td><td>Merck</td><td>1.05833</td><td>Concentration in Ringer solution: 0.8 mM</td></tr><tr><td>N[2-Hydroxyethyl]piperazine-N&#039;-[2-ethanesulfonic acid] (HEPES)</td><td>Formedium</td><td>HEPES10</td><td>Concentration in Ringer solution: 10 mM</td></tr><tr><td>Neo 5.5 sCMOS camera</td><td>ANDOR</td><td>DC-152Q-FI</td><td></td></tr><tr><td>NIS-Elements imaging software</td><td>Nikon</td><td>AR</td><td></td></tr><tr><td>Pluronic acid F-127</td><td>Millipore</td><td>540025</td><td></td></tr><tr><td>Pottasium chloride</td><td>Bio-Lab</td><td>163823</td><td>Concentration in Ringer solution: 5.4 mM</td></tr><tr><td>Pyrithione</td><td>Sigma Aldrich</td><td>H3261</td><td>Concentration in Ringer zinc solution: 7 &amp;mu;M</td></tr><tr><td>Silicone Grease Kit</td><td>Warner Instruments</td><td>W4 64-0378</td><td></td></tr><tr><td>Sodium chloride</td><td>Bio-Lab</td><td>190305</td><td>Concentration in Ringer solution: 120 mM</td></tr><tr><td>Zinc sulfate</td><td>Sigma Aldrich</td><td>31665</td><td>Concentration in Ringer zinc solution: 7 &amp;mu;M</td></tr></tbody></table>

characterizing mammalian zinc transporters using an in vitro zinc transport assay

Transition metals such as Zn2+ ions must be tightly regulated due to their cellular toxicity. Previously, the activity of Zn2+ transporters was measured indirectly by determining the expression level of the transporter under different concentrations of Zn2+. This was done by utilizing immunohistochemistry, measuring mRNA in the tissue, or determining the cellular Zn2+ levels. With the development of intracellular Zn2+ sensors, the activities of zinc transporters are currently primarily determined by correlating changes in intracellular Zn2+, detected using fluorescent probes, with the expression of the Zn2+ transporters. However, even today, only a few labs monitor dynamic changes in intracellular Zn2+ and use it to measure the activity of zinc transporters directly. Part of the problem is that out of the 10 zinc transporters of the ZnT family, except for ZnT10 (transports manganese), only zinc transporter 1 (ZnT1) is localized at the plasma membrane. Therefore, linking the transport activity to changes in the intracellular Zn2+ concentration is hard. This article describes a direct way to determine the zinc transport kinetics using an assay based on a zinc-specific fluorescent dye, FluoZin-3. This dye is loaded into mammalian cells in its ester form and then trapped in the cytosol due to cellular di-esterase activity. The cells are loaded with Zn2+ by utilizing the Zn2+ ionophore pyrithione. The ZnT1 activity is assessed from the linear part of the reduction in fluorescence following the cell washout. The fluorescence measured at an excitation of 470 nm and emission of 520 nm is proportional to the free intracellular Zn2+. Selecting the cells expressing ZnT1 tagged with the mCherry fluorophore allows for monitoring only the cells expressing the transporter. This assay is used to investigate the contribution of different domains of ZnT1 protein to the transport mechanism of human ZnT1, a eukaryotic transmembrane protein that extrudes excess zinc from the cell.

ZnT1 is a mammalian zinc transporter located on the cell plasma membrane13. It is a member of the cation diffusion facilitator (CDF) protein family that extrudes zinc from the cytosol to the extracellular millieu14. ZnT1 has a two-domain architecture: the transmembrane domain, which transports the ions across the membrane, and a C-terminal domain14. Unlike other known CDF proteins, ZnT1 has an extended unstructured C-terminal domain (USCTD). The role...

Watch this Scientific Journal Video about Exploring Cellular Zinc Regulation Through ZnT1 Functionality at JoVE.com

Author Spotlight: Exploring Cellular Zinc Regulation Through ZnT1 Functionality 

Zinc transport has proven challenging to measure due to the weak causal links to protein function and the low temporal resolution. This protocol describes a method for monitoring, with high temporal resolution, Zn2+ extrusion from living cells by utilizing a Zn2+ sensitive fluorescent dye, thus providing a direct measure of Zn2+ efflux.

Characterizing Mammalian Zinc Transporters Using an In Vitro Zinc Transport Assay

Transition metals such as Zn2+ ions must be tightly regulated due to their cellular toxicity. Previously, the activity of Zn2+ transporters was measured ...

characterizing-mammalian-zinc-transporters-using-an-vitro-zinc

Research

JoVE Journal

Biology

1.1K Views.  Ben-Gurion University. Mijn onderzoek richt zich op de structuur-functie relatie van zinktransporters bij zoogdieren, met speciale nadruk op het regulatiemechanisme. Vanwege het belang van zinkionen voor de cellulaire functie, is het een uitdaging om nauwkeurige metingen van de cellulaire zinkdynamiek te verkrijgen. Ons protocol richt zich op de lage signaal-ruisverhouding en lage temporele resolutie van zinkdynamica.Deze methode maakt het mogelijk om de instroom van zink door het celmembraan te testen, met ...

Video: Author Spotlight: Verkenning van cellulaire zinkregulatie door middel van ZnT1-functionaliteit 

Atomic Absorbance Spectroscopy to Measure Intracellular Zinc Pools in Mammalian Cells

Transition metals are essential micronutrients for organisms but can be toxic to cells at high concentrations by competing with physiological metals in proteins and generating redox stress. Pathological conditions that lead to metal depletion or accumulation are causal agents of different human diseases. Some examples include anemia, acrodermatitis enteropathica, and Wilson&#8217;s and Menkes&#8217; diseases. It is therefore important to be able to measure the levels and transport of transition metals in biological samples with high sensitivity and accuracy in order to facilitate research exploring how these elements contribute to normal physiological functions and toxicity. Zinc (Zn), for example, is a cofactor in many mammalian proteins, participates in signaling events, and is a secondary messenger in cells. In excess, Zn is toxic and can inhibit absorption of other metals, while in deficit, it can lead to a variety of potentially lethal conditions.

Graphite furnace atomic absorption spectroscopy (GF-AAS) provides a highly sensitive and effective method for determining Zn and other transition metal concentrations in diverse biological samples. Electrothermal atomization via GF-AAS quantifies metals by atomizing small volumes of samples for subsequent selective absorption analysis using wavelength of excitation of the element of interest. Within the limits of linearity of the Beer-Lambert Law, the absorbance of light by the metal is directly proportional to concentration of the analyte. Compared to other methods of determining Zn content, GF-AAS detects both free and complexed Zn in proteins and possibly in small intracellular molecules with high sensitivity in small sample volumes. Moreover, GF-AAS is also more readily accessible than inductively coupled plasma mass spectrometry (ICP-MS) or synchrotron-based X-ray fluorescence. In this method, the systematic sample preparation of different cultured cell lines for analyses in a GF-AAS is described. Variations in this trace element were compared in both whole cell lysates and subcellular fractions of proliferating and differentiated cells as proof of principle.

Cultured primary or established cell lines are commonly used to address fundamental biological and mechanistic questions as an initial approach before using animal models. This protocol describes how to prepare whole cell extracts and subcellular fractions for studies of zinc (Zn) and other trace elements with atomic absorbance spectroscopy.

Transition metals are essential micronutrients for organisms but can be toxic to cells at high concentrations by competing with physiological metals in ...

Biochemistry

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine exchangers in plants have not been identified. Here, we outline a method to characterize polyamine antiporters using membrane vesicles generated from the lysis of Escherichia coli cells heterologously expressing a plant antiporter. First, we heterologously expressed AtBAT1 in an E. coli strain deficient in polyamine and arginine exchange transporters. Vesicles were produced using a French press, purified by ultracentrifugation and utilized in a membrane filtration assay of labeled substrates to demonstrate the substrate specificity of the transporter. These assays demonstrated that AtBAT1 is a proton-mediated transporter of arginine, &#947;-aminobutyric acid (GABA), putrescine and spermidine. The mutant strain that was developed for the assay of AtBAT1 may be useful for the functional analysis of other families of plant and animal polyamine exchangers. We also hypothesize that this approach can be used to characterize many other types of antiporters, as long as these proteins can be expressed in the bacterial cell membrane. E. coli is a good system for the characterization of novel transporters, since there are multiple methods that can be employed to mutagenize native transporters.

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

We describe a method for the characterization of proton-driven membrane transporters in membrane vesicle preparations produced by heterologous expression in E. coli and lysis of cells using a French press.

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine ...

High-throughput Screening for Small-molecule Modulators of Inward Rectifier Potassium Channels

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, atrial fibrillation, and pain1,2. For the most part, however, progress toward understanding their therapeutic potential or even basic physiological functions has been slowed by the lack of good pharmacological tools. Indeed, the molecular pharmacology of the inward rectifier family has lagged far behind that of the S4 superfamily of voltage-gated potassium (Kv) channels, for which a number of nanomolar-affinity and highly selective peptide toxin modulators have been discovered3. The bee venom toxin tertiapin and its derivatives are potent inhibitors of Kir1.1 and Kir3 channels4,5, but peptides are of limited use therapeutically as well as experimentally due to their antigenic properties and poor bioavailability, metabolic stability and tissue penetrance. The development of potent and selective small-molecule probes with improved pharmacological properties will be a key to fully understanding the physiology and therapeutic potential of Kir channels.
The Molecular Libraries Probes Production Center Network (MLPCN) supported by the National Institutes of Health (NIH) Common Fund has created opportunities for academic scientists to initiate probe discovery campaigns for molecular targets and signaling pathways in need of better pharmacology6. The MLPCN provides researchers access to industry-scale screening centers and medicinal chemistry and informatics support to develop small-molecule probes to elucidate the function of genes and gene networks. The critical step in gaining entry to the MLPCN is the development of a robust target- or pathway-specific assay that is amenable for high-throughput screening (HTS).
Here, we describe how to develop a fluorescence-based thallium (Tl+) flux assay of Kir channel function for high-throughput compound screening7,8,9,10.The assay is based on the permeability of the K+ channel pore to the K+ congener Tl+. A commercially available fluorescent Tl+ reporter dye is used to detect transmembrane flux of Tl+ through the pore. There are at least three commercially available dyes that are suitable for Tl+ flux assays: BTC, FluoZin-2, and FluxOR7,8. This protocol describes assay development using FluoZin-2. Although originally developed and marketed as a zinc indicator, FluoZin-2 exhibits a robust and dose-dependent increase in fluorescence emission upon Tl+ binding. We began working with FluoZin-2 before FluxOR was available7,8 and have continued to do so9,10. However, the steps in assay development are essentially identical for all three dyes, and users should determine which dye is most appropriate for their specific needs. We also discuss the assay's performance benchmarks that must be reached to be considered for entry to the MLPCN. Since Tl+ readily permeates most K+ channels, the assay should be adaptable to most K+ channel targets.

Methods for developing and validating a quantitative fluorescence assay for measuring the activity of inward rectifier potassium (Kir) channels for high-throughput compound screening is presented.

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, ...

Metal-Limited Growth of Neisseria gonorrhoeae for Characterization of Metal-Responsive Genes and Metal Acquisition from Host Ligands

Trace metals such as iron and zinc are vital nutrients known to play key roles in prokaryotic processes including gene regulation, catalysis, and protein structure. Metal sequestration by hosts often leads to metal limitation for the bacterium. This limitation induces bacterial gene expression whose protein products allow bacteria to overcome their metal-limited environment. Characterization of such genes is challenging. Bacteria must be grown in meticulously prepared media that allows sufficient access to nutritional metals to permit bacterial growth while maintaining a metal profile conducive to achieving expression of the aforementioned genes. As such, a delicate balance must be established for the concentrations of these metals. Growing a nutritionally fastidious organism such as Neisseria gonorrhoeae, which has evolved to survive only in the human host, adds an additional level of complexity. Here, we describe the preparation of a defined metal-limited medium sufficient to allow gonococcal growth and the desired gene expression. This method allows the investigator to chelate iron and zinc from undesired sources while supplementing the media with defined sources of iron or zinc, whose preparation is also described. Finally, we outline three experiments that utilize this media to help characterize the protein products of metal-regulated gonococcal genes.

Metal-Limited Growth of Neisseria gonorrhoeae for Characterization of Metal-Responsive Genes and Metal Acquisition from Host Ligands

We describe here a method for growth of Neisseria gonorrhoeae in metal-restricted liquid medium to facilitate the expression of genes for metal uptake. We also outline downstream experiments to characterize the phenotype of gonococci grown in these conditions. These methods may be adapted to be suitable for characterization of metal-responsive genes in other bacteria.

Trace metals such as iron and zinc are vital nutrients known to play key roles in prokaryotic processes including gene regulation, catalysis, and protein ...

Immunology and Infection

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating gastric H+,K+-ATPase is more difficult to investigate. Many transport assays utilize radioisotopes to achieve a sufficient signal-to-noise ratio, however, the necessary security measures impose severe restrictions regarding human exposure or assay design. Furthermore, ion transport across cell membranes is critically influenced by the membrane potential, which is not straightforwardly controlled in cell culture or in proteoliposome preparations. Here, we make use of the outstanding sensitivity of atomic absorption spectrophotometry (AAS) towards trace amounts of chemical elements to measure Rb+ or Li+ transport by Na+,K+- or gastric H+,K+-ATPase in single cells. Using Xenopus oocytes as expression system, we determine the amount of Rb+ (Li+) transported into the cells by measuring samples of single-oocyte homogenates in an AAS device equipped with a transversely heated graphite atomizer (THGA) furnace, which is loaded from an autosampler. Since the background of unspecific Rb+ uptake into control oocytes or during application of ATPase-specific inhibitors is very small, it is possible to implement complex kinetic assay schemes involving a large number of experimental conditions simultaneously, or to compare the transport capacity and kinetics of site-specifically mutated transporters with high precision. Furthermore, since cation uptake is determined on single cells, the flux experiments can be carried out in combination with two-electrode voltage-clamping (TEVC) to achieve accurate control of the membrane potential and current. This allowed e.g. to quantitatively determine the 3Na+/2K+ transport stoichiometry of the Na+,K+-ATPase and enabled for the first time to investigate the voltage dependence of cation transport by the electroneutrally operating gastric H+,K+-ATPase. In principle, the assay is not limited to K+-transporting membrane proteins, but it may work equally well to address the activity of heavy or transition metal transporters, or uptake of chemical elements by endocytotic processes.

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

We describe a method to quantify the activity of K+-countertransporting P-type ATPases by heterologous expression of the enzymes in Xenopus oocytes and measuring Rb+ or Li+ uptake into individual cells by atomic absorption spectrophotometry. The method is a sensitive and safe alternative to radioisotope flux experiments facilitating complex kinetic studies.

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating ...

In vitro Investigation of the MexAB Efflux Pump From Pseudomonas aeruginosa

There is an emerging scientific need for reliable tools for monitoring membrane protein transport. We present a methodology leading to the reconstitution of efflux pumps from the Gram-negative bacteria Pseudomonas aeruginosa in a biomimetic environment that allows for an accurate investigation of their activity of transport. Three prerequisites are fulfilled: compartmentation in a lipidic environment, use of a relevant index for transport, and generation of a proton gradient. The membrane protein transporter is reconstituted into liposomes together with bacteriorhodopsin, a light-activated proton pump that generates a proton gradient that is robust as well as reversible and tunable. The activity of the protein is deduced from the pH variations occurring within the liposome, using pyranin, a pH-dependent fluorescent probe. We describe a step-by-step procedure where membrane protein purification, liposome formation, protein reconstitution, and transport analysis are addressed. Although they were specifically designed for an RND transporter, the described methods could potentially be adapted for use with any other membrane protein transporter energized by a proton gradient.

In vitro Investigation of the MexAB Efflux Pump From Pseudomonas aeruginosa

We present a protocol for the in vitro investigation of efflux pumps from Pseudomonas aeruginosa. This protocol allows for the generation of a robust, reversible, and tunable proton gradient within the liposome membrane and hence should be adaptable to any membrane protein energized by the protomotive force.

There is an emerging scientific need for reliable tools for monitoring membrane protein transport. We present a methodology leading to the reconstitution ...

A Proteoliposome-Based Efflux Assay to Determine Single-molecule Properties of Cl- Channels and Transporters

The last 15 years have been characterized by an explosion in the ability to overexpress and purify membrane proteins from prokaryotic organisms as well as from eukaryotes. This increase has been largely driven by the successful push to obtain structural information on membrane proteins. However, the ability to functionally interrogate these proteins has not advanced at the same rate and is often limited to qualitative assays of limited quantitative value, thereby limiting the mechanistic insights that they can provide. An assay to quantitatively investigate the transport activity of reconstituted Cl- channels or transporters is described. The assay is based on the measure of the efflux rate of Cl- from proteoliposomes following the addition of the K+ ionophore valinomycin to shunt the membrane potential. An ion sensitive electrode is used to follow the time-course of ion efflux from proteoliposomes reconstituted with the desired protein. The method is highly suited for mechanistic studies, as it allows for the quantitative determination of key properties of the reconstituted protein, such as its unitary transport rate, the fraction of active protein and the molecular mass of the functional unit. The assay can also be utilized to determine the effect of small molecule compounds that directly inhibit/activate the reconstituted protein, as well as to test the modulatory effects of the membrane composition or lipid-modifying reagents. Where possible, direct comparison between results obtained using this method were found to be in good agreement with those obtained using electrophysiological approaches. The technique is illustrated using CLC-ec1, a CLC-type H+/Cl- exchanger, as a model system. The efflux assay can be utilized to study any Cl- conducting channel/transporter and, with minimal changes, can be adapted to study any ion-transporting protein.

A Proteoliposome-Based Efflux Assay to Determine Single-molecule Properties of Cl- Channels and Transporters

Proteoliposomes are used to study purified channels and transporters reconstituted in a well-defined biochemical environment. An experimental procedure to measure efflux mediated by these proteins is illustrated. The steps to prepare proteoliposomes, perform the recordings, and analyze data to quantitatively determine the functional properties of the reconstituted protein are described.

The last 15 years have been characterized by an explosion in the ability to overexpress and purify membrane proteins from prokaryotic organisms as well as ...

Direct Protein Delivery to Mammalian Cells Using Cell-permeable Cys2-His2 Zinc-finger Domains

Due to their modularity and ability to be reprogrammed to recognize a wide range of DNA sequences, Cys2-His2 zinc-finger DNA-binding domains have emerged as useful tools for targeted genome engineering. Like many other DNA-binding proteins, zinc-fingers also possess the innate ability to cross cell membranes. We recently demonstrated that this intrinsic cell-permeability could be leveraged for intracellular protein delivery. Genetic fusion of zinc-finger motifs leads to efficient transport of protein and enzyme cargo into a broad range of mammalian cell types. Unlike other protein transduction technologies, delivery via zinc-finger domains does not inhibit enzyme activity and leads to high levels of cytosolic delivery. Here a detailed step-by-step protocol is presented for the implementation of zinc-finger technology for protein delivery into mammalian cells. Key steps for achieving high levels of intracellular zinc-finger-mediated delivery are highlighted and strategies for maximizing the performance of this system are discussed.

Direct Protein Delivery to Mammalian Cells Using Cell-permeable Cys2-His2 Zinc-finger Domains

Zinc-finger domains are intrinsically cell-permeable and capable of mediating protein delivery into a broad range of mammalian cell types. Here, a detailed step-by-step protocol for implementing zinc-finger technology for intracellular protein delivery is presented.

Due to their modularity and ability to be reprogrammed to recognize a wide range of DNA sequences, Cys2-His2 zinc-finger DNA-binding domains have emerged ...

Assessing Mineral Availability in Fish Feeds using Complementary Methods Demonstrated with the Example of Zinc in Atlantic Salmon

Assessing the availability of dietary micro-minerals is a major challenge in mineral nutrition of fish species. The present article aims to describe a systematic approach combining different methodologies to assess the availability of zinc (Zn) in Atlantic salmon (Salmo salar). Considering that several Zn chemical species can be present in an Atlantic salmon feed, it was hypothesised that Zn availability is influenced by the Zn chemical species present in the feed. Thus, in this study, the first protocol is about how to extract the different Zn chemical species from the feed and to analyze them by a size exclusion chromatography-inductively coupled plasma mass spectroscopy (SEC-ICP-MS) method. Subsequently, an in vitro method was developed to evaluate the solubility of dietary Zn in Atlantic salmon feeds. The third protocol describes the method to study the impact of changing Zn chemical species composition on the uptake of Zn in a fish intestinal epithelial model using a rainbow trout gut cell line (RTgutGC). Together, the findings from the in vitro methods were compared with an in vivo study examining the apparent availability of inorganic and organic sources of Zn supplemented to Atlantic salmon feeds. The results showed that several Zn chemical species can be found in feeds and the efficiency of an organic Zn source depends very much on the amino acid ligand used to chelate Zn. The findings of the in vitro methods had less correlation with that outcome of the in vivo study. Nevertheless, in vitro protocols described in this article provided crucial information regarding Zn availability and its assessment in fish feeds.

This article explains in detail a systematic approach to assess micro-mineral availability in Atlantic salmon. The methodology includes tools and models with increasing biological complexity: (1) chemical speciation analysis, (2) in vitro solubility, (3) uptake studies in cell lines, and (4) in vivo fish studies.&#160;

Assessing the availability of dietary micro-minerals is a major challenge in mineral nutrition of fish species. The present article aims to describe a ...

Assessing Bioindicator Phenotypes for Zinc Limitation in M. smegmatis

Using Mycobacterium smegmatis as a Bioindicator for Zinc-Limited Growth Conditions in Mycobacteria

Many bacteria build alternative ribosomes in Zn2+-limiting growth conditions by replacing Zn2+-binding ribosomal proteins with Zn2+-independent paralogs. Defining a system to study these alternative ribosomes has proven difficult because Zn2+ contamination in the laboratory is common. To address this issue, chelating agents are sometimes added to growth media, but this approach convolutes the biological response to gradual Zn2+&#160;limitation and is associated with ribosome hibernation. Here, detailed instructions are outlined for preparing media and seeding cultures for Zn2+-limited growth without adding chelators. Following this method, the model bacterium, Mycobacterium smegmatis, undergoes morphogenesis, which depends on alternative ribosomes. Because morphogenesis is tractable and only occurs in Zn2+-limiting conditions, M. smegmatis can be used as a bioindicator to verify biologically relevant growth conditions. Three bioindicator phenotypes (cell density, cell length, and coenzyme F420 fluorescence) that indicate Zn2+ limitation in the wild-type are described, and changes in these bioindicators for a deletion mutant that cannot build alternative ribosomes are outlined. Since trace Zn2+ contamination is difficult to control for each batch of media, and precise quantification of Zn2+ in each media preparation is overly burdensome, following this bioindicator phenotype is an accessible way to validate the preparation of Zn2+-limited growth media. To help identify proper conditions for Zn2+-limiting growth and alternative ribosome production, changes in the bioindicator phenotypes were profiled for Zn2+-contaminated or severely Zn2+-depleted preparations of Zn2+-limited media as well. Further details to achieve Zn2+-limiting growth and alternative ribosome production in M. tuberculosis are presented, along with the associated bioindicator phenotype. Overall, the detailed instructions and bioindicator phenotypes described here will help standardize the production of translationally active alternative ribosomes in mycobacteria.

This protocol details preparation of Zn2+-limited media, growth vessels, and seed cultures to produce Zn2+-limited ribosomes in mycobacteria. Traces of Zn2+ lead to Zn2+-limiting growth and we describe morphogenic features used as bioindicators to verify Zn2+ limitation in vitro in the model bacterium, Mycobacterium smegmatis, and the human pathogen M. tuberculosis.

Many bacteria build alternative ribosomes in Zn2+-limiting growth conditions by replacing Zn2+-binding ribosomal proteins with Zn2+-independent paralogs. ...

Characterizing Mammalian Zinc Transporters Using an In Vitro Zinc Transport Assay

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