JMD and MKB are funded through the IGERT Nanomedicine Science and Technology program at Northeastern University (funding from NCI and NSF grant DGE- 0504331). This work was also funded by National Institutes of Health National Institute of General Medical Sciences Grant R01 GM084366. We also thank Saumya Das and Anthony Rosenzweig of BIDMC in Boston for providing cardiac myocytes and Kevin Cash for his contribution in the animal protocol development.

1. Preparation of optode
Before making the optode, aliquots of the components are need so that they can be easily measured and stored.
<ol>
	<li>A 50 mg Sodium Ionophore X (NaIX) vial and a 50 mg sodium tetrakis[3,5-bis(trifluoromethyl)phenyl]borate (NaTFPB) will each be brought up in tetrahyrdofuran (THF) and aliquoted into 1.5 ml polystyrene centrifuge tubes. In a chemical fume hood, dissolve the 50 mg of NaIX in 1 ml of THF in the shipping vial and mix to ensure that the solid has completely dissolved. Transfer 100 &mu;l of this solution into 10 separate centrifuge tubes. Repeat for NaTFPB. Allow the THF to evaporate in a chemical fume hood overnight, label the centrifuge tubes and place them in a 4 degree refrigerator for storage. This creates 5 mg aliquots of dry NaIX and NaTFPB.</li>
	<li>Dissolve the Chromoionophore III (CHIII) in 1 ml of THF and transfer to a 3 ml glass vial. Add another 1 ml of THF to the CHIII shipping vial to dissolve any residual CHIII and add this to the 3 ml glass vial. There will be a total of 2 ml now. Transfer 1 ml of the CHIII in THF solution to two separate 1.5 ml glass vials with a Teflon coated caps. Store the CHIII solution in a 4 degree refrigerator at the final concentration of 5 mg/ml. </li>
	</ol>
	These aliquots and solutions will be used to make the optode material.
	<ol start="3">
	<li>Pipette 66 &mu;l of bis(2-ethylhexyl)sebacate (DOS) into a 1.5 ml glass vial with a Teflon coated screw cap - this is the optode vial. DOS is a viscous solution so care must be taken to ensure the proper amount of solution is transferred to the vial. The volume corresponds to &tilde; 60 mg of DOS. </li>
	<li> Weigh out 30 mg of high molecular weight poly(vinyl) chloride (PVC) and transfer this to the optode vial.</li>
	<li>Now add the aliquoted functional components to the optode vial to create the desired optode. The relative concentrations of CHIII, NaIX, and NaTFPB will determine the response of the sensor to sodium. These concentrations can be adjusted to have a sensor that ideally responds to intracellular concentrations, with a Kd of 10 mM, or extracellular concentrations, with a Kd of 150 mM. Additionally, there will likely be batch to batch variability of the chemicals from the vendor and calibration of the sensors is necessary to determine if the correct response is occurring for each new batch of chemical components. Here we describe the method to create a sensor with concentrations that are typically used for intracellular sodium measurements. </li>
	<li>In a chemical fume hood, transfer 100 &mu;l of the 5 mg/ml CHIII in THF solution to the optode vial. </li>
	<li> Dissolve 5 mg of NaIX in 300 &mu;l of THF and transfer 300 &mu;l to the optode vial, this correlates to 5 mg of NaIX. </li>
	<li>Dissolve a 5 mg aliquot of the NaTFPB in 500 &mu;l of THF and transfer 20 &mu;l of the solution to the optode vial, this correlates to 0.1 mg of NaTFPB. </li>
	<li>Add 80 &mu;l of THF to the optode vial.</li>
	<li>The solution in the optode vial contains 30 mg of PVC, 60 mg of DOS, 5 mg of NaIX, 0.2 mg of NaTFPB, 0.5 mg of CHIII in 500 &mu;l of THF. Gently vortex this vial until all of the PVC has dissolved. The optode should be a red color. The total amount of THF added to the vial should be 500 &mu;L regardless of the ratio of components used. </li>
	<li>Allow the remaining THF in the NaIX and NaTFPB aliquots to evaporate overnight and re-label the tube with the correct amount of chemical.</li>
</ol>
2. Creating nanosensors
<ol>
	<li>Pipette 100 &mu;l of 10 mg/ml 1,2-Distearoyl-sn-Glycero-3-Phosphoethanolamine-N -[Methoxy(Polyethyleneglycol)-550] in chloroform (PEG-lipid) into a 4 dram glass vial. Allow the chloroform to evaporate in a chemical fume hood. This process can be expedited by drying the solution with nitrogen gas or house air. </li>
	<li>Add 4 ml of the desired aqueous solution to the vial with PEG-lipid. Place the vial on a laboratory jack underneath the sonicating tip and raise the vial until the tip is about 7 mm deep in the solution and sonicate at a low sonication power for 30 seconds. Here we use a Branson digital sonifier set to 15% amplitude with a &frac12; inch tip diameter step horn. The aqueous solution can be any solution. For example, to determine the response of the sensors we use 10 mM HEPES pH 7.2 with trizma base. </li>
	<li>While the solution is being sonicated, mix 50 &mu;l of the optode material with 50 &mu;l of dichloromethane in a 0.6 ml microcentrifuge tube. Mix with the pipette to ensure even mixing.</li>
	<li>Adjust the sonifier control to sonicate for 3 minutes. Start the sonication and while sonicating add the optode mixture solution to the vial by immersing the pipette tip into the solution and dispensing the 100 &mu;l of solution in one quick, even injection. The solution should turn blue, depending on the aqueous solution used. </li>
	<li>Allow the sonication to continue for about 30 seconds then lower the jack so that the sonicating tip is about 2 mm deep in solution. This should begin to aerate the solution and the solution will become opaque. This step helps to remove the residual THF and dichloromethane from the solution. </li>
	<li>Once sonication is complete, pull up the nanosensor solution into a 5 ml syringe. Attach a 0.2 &mu;m syringe filter and dispense the nanosensor solution into a glass vial with screw top cap. The solution should be clear and blue if an aqueous solution that contains less than 10 mM sodium and has a pH less than about 9 is used. Otherwise, it should have no color or be pink.</li>
</ol>
3. Determining nanonsensor response
This method is used to determine the response of nanosensors designed to measure intracellular sodium. Different concentrations are recommended to be used for extracellular sodium concentration nanosensors. 
<ol>
	<li>Make stock solutions of 10 mM HEPES (0 Na) and 1M sodium chloride (NaCl) in 10 mM HEPES (1 M Na) and pH each of these solutions to 7.2 using trizma base. Mix stock solutions of 0 Na and 1 M Na in 50 ml conical centrifuge tubes to create solutions with: 0, 1, 5, 10, 20, 50, 100, 200, 500, 1000 mM NaCl concentrations. </li>
	<li>Use an optical bottom 96 well plate to determine the response. Add 100 &mu;l of each concentration of sodium to 3 wells in a column. This will produce an array of 3 x 10 wells.</li>
	<li>To each well add 100 &mu;l of freshly prepared nanosensors and load into a microplate fluorometer. We use a Molecular Devices Spectramax M3.</li>
	<li>Read each well with the following wavelengths: </li>
</ol>
<table border="1">
	<tbody>
		<tr>
			<td>Excitation (nm)</td>
			<td>Emission (nm)</td>
			<td>Cut-Off (nm)</td>
		</tr>
		<tr>
			<td>488</td>
			<td>570</td>
			<td>530</td>
		</tr>
		<tr>
			<td>488</td>
			<td>670</td>
			<td>610</td>
		</tr>
		<tr>
			<td>639</td>
			<td>680</td>
			<td>665</td>
		</tr>
	</tbody>
</table>
 The nanosensor intensity at each wavelength is dependent on the sodium concentration. The use of the wavelength depends on the applications. For intracellular slow dynamics measurements, we use 488 nm/570 nm (excitation/emission) and 488 nm/670 nm which allow us to ratio the two wavelengths and reduce noise. 
<ol start="5">
<li>Traditionally, optode data is converted to a values which are: &alpha; = (I - Imin)/(Imax - Imin), where I is the intensity at a given sodium concentration, Imin is the lowest intensity in the response measurements, and Imax is the highest intensity in the response measurements. Convert either the ratio of intensity at 570 nm divided by the intensity at 670 nm or the intensity at 680 nm to &alpha;. </li>
<li>Use plotting software, either Excel or Origin are typically used in our lab, to plot &alpha; versus the log of the sodium concentration, setting the log of zero sodium concentration to -1. Fit a sigmoidal curve to the response. From the curve, calculate the sodium concentration at &alpha; = 0.5 which represents the concentration at which the sensor optimally responds. </li>
<li>Adjust the ratio of CHIII, NaIX, and NaTFPB in the optode to create nanosensors with optimal response around the sodium concentration of interest. </li>
</ol>
4. Intracellular imaging
<ol>
	<li>For intracellular measurements, the nanosensors are made in 5 mg/ml glucose in water and microinjected into the cells. </li>
	<li>Grow or transfer cells to a glass bottom dish or an imaging chamber with a glass coverslip bottom. </li>
	<li>Incubate the cells in clear media or Tyrode's solution. Mount the imaging chamber or dish onto an epifluorescence microscope. We use a Zeiss LSM 7 confocal microscope. </li>
	<li>Pull capillary glass with a pipette puller. We use a sutter flaming brown p-97 with 1 mm OD, 0.78 mm ID glass, to a final tip diameter around 300-500 nm. </li>
	<li>Backfill the pipette with nanosensor solution using a syringe and a Hamilton needle, or a microtip loader and a pipette.</li>
	<li>Mount the pipette into a holder that is attached to a micromanipulator and connected to a pressure controlled injection system. Here we use a Burleigh EXFO PCS 6000 micromanipulator and a medical systems corporation injector.</li>
	<li>Lower the pipette on top of the cell and into the cytoplasm, sometimes it is necessary to "tap" the manipulator holder to pierce the membrane. Inject the nanosensor solution and quickly withdraw the pipette. </li>
</ol>
5. In vivo imaging
All procedures have been reviewed and approved by Northeastern University's Animal Care and Use Committee.
<ol>
	<li>Nanosensors for in vivo, extracellular imaging are made in phosphate buffered saline and adjusted to have an optimal response to sodium at a sodium concentration of 135 mM.</li>
	<li>For the injection set-up and imaging of the fluorescent nanosensors in mice, we use the IVIS Lumina II and its associated anesthesia unit. Disinfect the induction and imaging chambers. Place CD1 nude mice (approximately 20g) in the induction chamber and expose them to isoflurane. The percent of isoflurane and oxygen flowrate administered will vary depending on the species and weight of mice. Wait for the mice to become fully anesthetized.</li>
	<li>After the mice are anesthetized, remove the mice from the induction chamber and place into the imaging chamber. Keep the mice under anesthesia. For each mouse, disinfect the desired injection areas.</li>
	<li>Fill a sterile 31G insulin syringe with 10 &mu;L of nanosensors making sure to remove all air bubbles.</li>
	<li>Using forceps, grasp a small fold of skin along the back of the mouse at the desired injection site. While grasping the skin, insert the syringe into the skin with the bevel facing up and the needle parallel to the skin.</li>
	<li>Prior to injecting the sensors, pull back on the syringe plunger to ensure the needle is not inserted into a blood vessel. Then, gently move the needle right and left to create a pocket within the skin. This will minimize the back pressure which causes the sensors to leak out of the injection site. Inject the sensors and gently remove the syringe.</li>
	<li>Gently apply pressure to the injection site and wipe away any solution that may have leaked out of the injection site. This will minimize fluorescence from sensors that are not injected into the skin.</li>
	<li>Repeats steps 5.4 through 5.7 until the number of desired injection areas is reached. Six injection sites evenly spaced on the left and right side of the back are recommended. For each individual mouse, change needles if the needle becomes difficult to insert into the skin. Needles should not be shared between mice. This will minimize disease transmission or cross-contamination amongst mice.</li>
	<li> For imaging the sodium fluorescent sensors, we acquire both brightfield and fluorescent images of the mice using filter sets that most closely matched the 640 nm/680 nm (excitation/emission) spectrum of the nanosensors and that also minimize the autofluorescence of the skin. </li>
</ol>
6. Representative Results
<img src="/files/ftp_upload/2896/2896fig1.jpg" alt="Figure 1"/> Figure 1. Response curves of five different nanosensor sets to sodium. Each set represents nanosensors from different optode formulations that had the same amount of CHIII, NaTFPB, PVC and DOS but varying amounts of NaIX . The data were converted to &alpha; values using the 639/680 nm intensities. Data are an average of 3, error bars omitted for clarity, with a fitted sigmoidal curve using Origin. In this response curve, the maximum concentration of sodium used was 500 mM. The Kd of the sodium nanosensors for sodium ranged from 10 mM (orange diamonds) to about 150 mM (black squares).
<img src="/files/ftp_upload/2896/2896fig2.jpg" alt="Figure 2"/> Figure 2. Injected neonatal cardiac myocytes. Cells were cultured on coverglass, mounted on a microscope and injected with sodium nanosensors. Shown are fluorescent (639 nm excitation), brightfield, and overlay. Note that some sensor clustering occurs but the cells have normal morphology, sensors have diffused throughout the cytosol and there is no nuclear loading. 
<img src="/files/ftp_upload/2896/2896fig3.jpg" alt="Figure 3"/> Figure 3. Subcutaneous injection of mice with sodium nanosensors. Nine different injections of sodium nanosensors were made into the subcutaneous space of nude mice. Both bright field and fluorescent images (640/680) are overlaid.

<ol>
	<li>Adrogue, H. J., Madias, N. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hyponatremia.">Hyponatremia.</a> N Engl J Med. 342, 1581-1589 (2000).</li><li>Kim, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BACE1+regulates+voltage-gated+sodium+channels+and+neuronal+activity.">BACE1 regulates voltage-gated sodium channels and neuronal activity.</a> Nat Cell Biol. 9, 755-764 (2007).</li><li>Dubach, J. M., Lim, E., Zhang, N., Francis, K. P., Clark, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=in+vivo+sodium+concentration+continuously+monitored+with+fluorescent+sensors.">in vivo sodium concentration continuously monitored with fluorescent sensors.</a> Integr Biol (Camb). , (2010).</li><li>Minta, A., Tsien, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+indicators+for+cytosolic+sodium.">Fluorescent indicators for cytosolic sodium.</a> J Biol Chem. 264, 19449-19457 (1989).</li><li>Dubach, J. M., Harjes, D. I., Clark, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ion-selective+nano-optodes+incorporating+quantum+dots.">Ion-selective nano-optodes incorporating quantum dots.</a> Journal of the American Chemical Society. 129, 8418-8418 (2007).</li><li>Dubach, J. M., Harjes, D. I., Clark, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+ion-selective+nanosensors+for+intracellular+analysis+with+improved+lifetime+and+size.">Fluorescent ion-selective nanosensors for intracellular analysis with improved lifetime and size.</a> Nano Letters. 7, 1827-1831 (1021).</li><li>Schaffar, B., Wolfbeis, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+sodium-selective+optrode.">A sodium-selective optrode.</a> Mikrochim Acta III. III, 109-109 (1989).</li><li>Seiler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+sodium-selective+optode+membranes+based+on+neutral+ionophores+and+assay+of+sodium+in+plasma.">Characterization of sodium-selective optode membranes based on neutral ionophores and assay of sodium in plasma.</a> Clin Chem. 37, 1350-1355 (1991).</li><li>Zhujun, Z., Mullin, J., Seitz, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+sensor+for+sodium+based+on+ion-pair+extraction+and+fluorescence.">Optical sensor for sodium based on ion-pair extraction and fluorescence.</a> Anal Chim Acta. 184, 251-251 (1986).</li><li>Bakker, E., Buhlmann, P., Pretsch, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Carrier-Based+Ion-Selective+Electrodes+and+Bulk+Optodes.+1.+General+Characteristics.">Carrier-Based Ion-Selective Electrodes and Bulk Optodes. 1. General Characteristics.</a> Chem Rev. 97, 3083-3132 (1997).</li><li>Dubach, J. M., Das, S., Rosenzweig, A., Clark, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+sodium+dynamics+in+isolated+cardiomyocytes+using+fluorescent+nanosensors.">Visualizing sodium dynamics in isolated cardiomyocytes using fluorescent nanosensors.</a> Proceedings of the National Academy of Sciences of the United States of America. 106, 16145-16150 (2009).</li><li>Harjes, D. I., Dubach, J. M., Rosenzweig, A., Das, S., Clark, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ion-Selective+Optodes+Measure+Extracellular+Potassium+Flux+in+Excitable+Cells.">Ion-Selective Optodes Measure Extracellular Potassium Flux in Excitable Cells.</a> Macromolecular Rapid Communications. 31, 217-221 (2010).</li><li>Balaconis, M. K., Billingsley, K. L., Dubach, J. M., Clark, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> , .</li><li>Park, E. J., Brasuel, M., Behrend, C., Philbert, M. 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No conflicts of interest declared.

The formation of nanosensors should take no longer than 10 minutes once the optode solution has been made and the PEG lipid dried. Optode can not only be made quickly when needed, but when stored at 4 degrees Celsius, they can be stable for months. We have shown that the nansensors, once formed, are stable in solution for at least a week; however, care must be taken to prevent photobleaching over this time by blocking the nanosensors from light.6 While sodium nanosensors were demonstrated here, optodes have been created for most biologically relevant ions such as potassium and chloride.10,12 We have also extended this technology to small molecules such as glucose.13
In addition to generating nanosensors to monitor different analytes, these nanosensors are amenable to other changes that will make them useful in most biological experiments. For example, the surface coating, in this case poly(ethylene glycol), can easily be changed using any amphiphilic molecule that is water soluble. This enables the possibility of functionalizing these nanoparticles for different applications or targets. The size of the nanoparticles can also be adjusted by altering the surface coating, the sonication intensity or the solvent used. 
Calcium fluorescent indicators have been invaluable in determining intracellular calcium signaling; however, no available sodium sensitive dyes have the same ideal characteristics. Optode nanoparticles intended to provide an alternative method for imaging ions intracellularly have been in development for years.14 We have built upon this previous research to create nanosensors specific for intracellular sodium imaging that will hopefully provide a means to understand how sodium signaling affects cellular function and alterations in signaling which lead to certain diseases.
The use of fluorescent nanosensors in vivo offers a real-time minimally-invasive alternative for monitoring analytes over other methods such as blood draws.3 Sodium and glucose nanosensors based on the technology above have been shown to track changes in sodium and glucose, respectively in vivo.3,13 However, there currently exist limitations to this method as a monitoring tool. For example, the sensors must contain a fluorophore with a spectrum sufficiently shifted towards the near infrared red in order to minimize background autofluorescence from the skin.15 Second, the current injection technique does not produce uniform injection of the nanosensors which may be caused by such errors as variation in injection depth. Despite these limitations, the development of these fluorescent nanosensors and their successful demonstration could make these sensors an invaluable research tool and method for monitoring patient health.

<table border="1">
	<tbody>
		<tr>
			<th>
				Product Name</th>
			<th>
				Company</th>
			<th>
				Catalogue Number</th>
			<th>
				Comments</th>
		</tr>
		<tr>
			<td>IVIS Lumina II Instrument Package</td>
			<td>Caliper Life Sciences</td>
			<td>126273</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CD-1 Nude Mice</td>
			<td>Charles River</td>
			<td>Strain Code: 086</td>
			<td>Immunodeficient</td>
		</tr>
		<tr>
			<td>Insulin syringes</td>
			<td>BD</td>
			<td>328438</td>
			<td>3/10 ccmL , 8mm, 31G</td>
		</tr>
		<tr>
			<td>High Molecular Weight PVC</td>
			<td>Sigma</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>DOS</td>
			<td>Sigma</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CHIII</td>
			<td>Sigma</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>NaTFPB</td>
			<td>Sigma</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>NaIX</td>
			<td>Sigma</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Thin walled capillary glass</td>
			<td>Sutter</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>PEG lipid</td>
			<td>Avanti Polar Lipids</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>
Table 1. Provides a list of chemicals and equipment used in this procedure. All common chemicals not listed were purchased from Sigma.

fluorescent nanoparticles for the measurement of ion concentration in biological systems

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion fluxes at the cellular level are required for initiating action potentials in excitable cells.2 Sodium regulation plays an important role in both of these cases; however, no method currently exists for continuously monitoring sodium levels in vivo 3 and intracellular sodium probes 4 do not provide similar detailed results as calcium probes. In an effort to fill both of these voids, fluorescent nanosensors have been developed that can monitor sodium concentrations in vitro and in vivo.5,6 These sensors are based on ion-selective optode technology and consist of plasticized polymeric particles in which sodium specific recognition elements, pH-sensitive fluorophores, and additives are embedded.7-9 Mechanistically, the sodium recognition element extracts sodium into the sensor. 10 This extraction causes the pH-sensitive fluorophore to release a hydrogen ion to maintain charge neutrality within the sensor which causes a change in fluorescence. The sodium sensors are reversible and selective for sodium over potassium even at high intracellular concentrations.6 They are approximately 120 nm in diameter and are coated with polyethylene glycol to impart biocompatibility. Using microinjection techniques, the sensors can be delivered into the cytoplasm of cells where they have been shown to monitor the temporal and spatial sodium dynamics of beating cardiac myocytes.11 Additionally, they have also tracked real-time changes in sodium concentrations in vivo when injected subcutaneously into mice.3 Herein, we explain in detail and demonstrate the methodology for fabricating fluorescent sodium nanosensors and briefly demonstrate the biological applications our lab uses the nanosensors for: the microinjection of the sensors into cells; and the subcutaneous injection of the sensors into mice.

Watch this Scientific Journal Video about Fluorescent Nanoparticles for the Measurement of Ion Concentration in Biological Systems at JoVE.com

Fluorescent Nanoparticles for the Measurement of Ion Concentration in Biological Systems

Fluorescent nanoparticles produced in our lab are used for imaging ion concentrations and ion fluxes in biological systems such as cells during signaling and interstitial fluid during physiological homeostasis.

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion ...

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15.2K Views.  Northeastern University. Fluorescent nanoparticles produced in our lab are used for imaging ion concentrations and ion fluxes in biological systems such as cells during signaling and interstitial fluid during physiological homeostasis.

Video: Fluorescent Nanoparticles for the Measurement of Ion Concentration in Biological Systems

Magnetic Resonance Elastography Methodology for the Evaluation of Tissue Engineered Construct Growth

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use of destructive assessment is not acceptable. A proposed alternative is the use of an imaging process called magnetic resonance elastography. Elastography is a nondestructive method for determining the engineered outcome by measuring local mechanical property values (i.e., complex shear modulus), which are essential markers for identifying the structure and functionality of a tissue. As a noninvasive means for evaluation, the monitoring of engineered constructs with imaging modalities such as magnetic resonance imaging (MRI) has seen increasing interest in the past decade1. For example, the magnetic resonance (MR) techniques of diffusion and relaxometry have been able to characterize the changes in chemical and physical properties during engineered tissue development2. The method proposed in the following protocol uses microscopic magnetic resonance elastography (&mu;MRE) as a noninvasive MR based technique for measuring the mechanical properties of small soft tissues3. MRE is achieved by coupling a sonic mechanical actuator with the tissue of interest and recording the shear wave propagation with an MR scanner4. Recently, &mu;MRE has been applied in tissue engineering to acquire essential growth information that is traditionally measured using destructive mechanical macroscopic techniques5. In the following procedure, elastography is achieved through the imaging of engineered constructs with a modified Hahn spin-echo sequence coupled with a mechanical actuator. As shown in Figure 1, the modified sequence synchronizes image acquisition with the transmission of external shear waves; subsequently, the motion is sensitized through the use of oscillating bipolar pairs. Following collection of images with positive and negative motion sensitization, complex division of the data produce a shear wave image. Then, the image is assessed using an inversion algorithm to generate a shear stiffness map6. The resulting measurements at each voxel have been shown to strongly correlate (R2&gt;0.9914) with data collected using dynamic mechanical analysis7. In this study, elastography is integrated into the tissue development process for monitoring human mesenchymal stem cell (hMSC) differentiation into adipogenic and osteogenic constructs as shown in Figure 2.

The procedure demonstrates the methodology of magnetic resonance elastography for monitoring the engineered outcome of adipose and osteogenic tissue engineered constructs through noninvasive local assessment of the mechanical properties using microscopic magnetic resonance elastography (&mu;MRE).

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use ...

Solid Lipid Nanoparticles (SLNs) for Intracellular Targeting Applications

Nanoparticle-based delivery vehicles have shown great promise for intracellular targeting applications, providing a mechanism to specifically alter cellular signaling and gene expression. In a previous investigation, the synthesis of ultra-small solid lipid nanoparticles (SLNs) for topical drug delivery and biomarker detection applications was demonstrated. SLNs are a well-studied example of a nanoparticle delivery system that has emerged as a promising drug delivery vehicle. In this study, SLNs were loaded with a fluorescent dye and used as a model to investigate particle-cell interactions. The phase inversion temperature (PIT) method was used for the synthesis of ultra-small populations of biocompatible nanoparticles. A 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenylphenyltetrazolium bromide (MTT) assay was utilized in order to establish appropriate dosing levels prior to the nanoparticle-cell interaction studies. Furthermore, primary human dermal fibroblasts and mouse dendritic cells were exposed to dye-loaded SLN over time and the interactions with respect to toxicity and particle uptake were characterized using fluorescence microscopy and flow cytometry. This study demonstrated that ultra-small SLNs, as a nanoparticle delivery system, are suitable for intracellular targeting of different cell types.

Solid Lipid Nanoparticles SLNs for Intracellular Targeting Applications

In this study, a method for synthesizing ultra-small populations of biocompatible nanoparticles was described, as well as several in vitro methods by which to assess their cellular interactions.

Nanoparticle-based delivery vehicles have shown great promise for intracellular targeting applications, providing a mechanism to specifically alter ...

Studying Soft-matter and Biological Systems over a Wide Length-scale from Nanometer and Micrometer Sizes at the Small-angle Neutron Diffractometer KWS-2

The KWS-2 SANS diffractometer is dedicated to the investigation of soft matter and biophysical systems covering a wide length scale, from nm to &#181;m. The instrument is optimized for the exploration of the wide momentum transfer Q range between 1x10-4 and 0.5 &#197;-1 by combining classical pinhole, focusing (with lenses), and time-of-flight (with chopper) methods, while simultaneously providing high-neutron intensities with an adjustable resolution. Because of its ability to adjust the intensity and the resolution within wide limits during the experiment, combined with the possibility to equip specific sample environments and ancillary devices, the KWS-2 shows a high versatility in addressing the broad range of structural and morphological studies in the field. Equilibrium structures can be studied in static measurements, while dynamic and kinetic processes can be investigated over time scales between minutes to tens of milliseconds with time-resolved approaches. Typical systems that are investigated with the KWS-2 cover the range from complex, hierarchical systems that exhibit multiple structural levels (e.g., gels, networks, or macro-aggregates) to small and poorly-scattering systems (e.g., single polymers or proteins in solution). The recent upgrade of the detection system, which enables the detection of count rates in the MHz range, opens new opportunities to study even very small biological morphologies in buffer solution with weak scattering signals close to the buffer scattering level at high Q.
In this paper, we provide a protocol to investigate samples with characteristic size levels spanning a wide length scale and exhibiting ordering in the mesoscale structure using KWS-2. We present in detail how to use the multiple working modes that are offered by the instrument and the level of performance that is achieved.

Here, we present a protocol to investigate soft matter and biophysical systems over a wide mesoscopic length scale, from nm to &#181;m that involves the use of the KWS-2 SANS diffractometer at high intensities and an adjustable resolution.

The KWS-2 SANS diffractometer is dedicated to the investigation of soft matter and biophysical systems covering a wide length scale, from nm to &#181;m. ...

Merging Ion Concentration Polarization between Juxtaposed Ion Exchange Membranes to Block the Propagation of the Polarization Zone

The ion concentration polarization (ICP) phenomenon is one of the most prevailing methods to preconcentrate low-abundance biological samples. The ICP induces a noninvasive region for charged biomolecules (i.e., the ion depletion zone), and targets can be preconcentrated on this region boundary. Despite the high preconcentration performances with ICP, it is difficult to find the operating conditions of non-propagating ion depletion zones. To overcome this narrow operating window, we recently developed a new platform for spatiotemporally fixed preconcentration. Unlike preceding methods that only use ion depletion, this platform also uses the opposite polarity of the ICP (i.e., ion enrichment) to stop the propagation of the ion depletion zone. By confronting the enrichment zone with the depletion zone, the two zones merge together and stop. In this paper, we describe a detailed experimental protocol to build this spatiotemporally defined ICP platform and characterize the preconcentration dynamics of the new platform by comparing them with those of the conventional device. Qualitative ion concentration profiles and current-time responses successfully capture the different dynamics between the merged ICP and the stand-alone ICP. In contrast to the conventional one that can fix the preconcentration location at only ~5 V, the new platform can produce a target-condensed plug at a specific location in the broad ranges of operating conditions: voltage (0.5-100 V), ionic strength (1-100 mM), and pH (3.7-10.3).

The protocol for a novel ion concentration polarization (ICP) platform that can stop the propagation of the ICP zone, regardless of the operating conditions is described. This unique ability of the platform lies in the use of merging ion depletion and enrichment, which are two polarities of the ICP phenomenon.

The ion concentration polarization (ICP) phenomenon is one of the most prevailing methods to preconcentrate low-abundance biological samples. The ICP ...

Production of Metal Nanoparticles by Pulsed Laser-ablation in Liquids: A Tool for Studying the Antibacterial Properties of Nanoparticles

The emergence of multidrug-resistant bacteria is a global clinical concern leading some to speculate about our return to a "pre-antibiotics" era of medicine. In addition to efforts to identify novel small-molecule antimicrobial drugs, there has been great interest in the use of metal nanoparticles as coatings for medical devices, wound dressings, and consumer packaging, due to their antimicrobial properties. The wide variety of methods available for nanoparticle synthesis results in a broad spectrum of chemical and physical properties which can affect antibacterial efficacy. This manuscript describes the pulsed laser-ablation in liquids (PLAL) method to create nanoparticles. This approach allows for the fine tuning of nanoparticle size, composition, and stability using post-irradiation methods as well as the addition of surfactants or volume excluders. By controlling particle size and composition, a large range of physical and chemical properties of metal nanoparticles can be explored which may contribute to their antimicrobial efficacy thereby opening new avenues for antibacterial development.

The antimicrobial properties of metals such as copper and silver have been recognized for centuries. This protocol describes pulsed laser-ablation in liquids, a method of synthesizing metal nanoparticles that provides the ability to fine tune the properties of these nanoparticles to optimize their antimicrobial effects.

The emergence of multidrug-resistant bacteria is a global clinical concern leading some to speculate about our return to a "pre-antibiotics" era of ...

Flash NanoPrecipitation for the Encapsulation of Hydrophobic and Hydrophilic Compounds in Polymeric Nanoparticles

The formulation of a therapeutic compound into nanoparticles (NPs) can impart unique properties. For poorly water-soluble drugs, NP formulations can improve bioavailability and modify drug distribution within the body. For hydrophilic drugs like peptides or proteins, encapsulation within NPs can also provide protection from natural clearance mechanisms. There are few techniques for the production of polymeric NPs that are scalable. Flash NanoPrecipitation (FNP) is a process that uses engineered mixing geometries to produce NPs with narrow size distributions and tunable sizes between 30 and 400 nm. This protocol provides instructions on the laboratory-scale production of core-shell polymeric nanoparticles of a target size using FNP. The protocol can be implemented to encapsulate either hydrophilic or hydrophobic compounds with only minor modifications. The technique can be readily employed in the laboratory at milligram scale to screen formulations. Lead hits can directly be scaled up to gram- and kilogram-scale. As a continuous process, scale-up involves longer mixing process run time rather than translation to new process vessels. NPs produced by FNP are highly loaded with therapeutic, feature a dense stabilizing polymer brush, and have a size reproducibility of &#177; 6%.

Flash NanoPrecipitation (FNP) is a scalable approach to produce polymeric core-shell nanoparticles. Lab-scale formulations for the encapsulation of hydrophobic or hydrophilic therapeutics are described.

The formulation of a therapeutic compound into nanoparticles (NPs) can impart unique properties. For poorly water-soluble drugs, NP formulations can ...

Continuous Measurement of Biological Noise in Escherichia Coli Using Time-lapse Microscopy

The protocol developed here offers a tool to enable computer tracking of Escherichia coli division and fluorescent levels over several hours. The process starts by screening for colonies that survive on minimal media, assuming that only Escherichia coli harboring the correct plasmid will be able to thrive in the specific conditions. Since the process of building large genetic circuits, requiring the assembly of many DNA parts, is challenging, circuit components are often distributed between multiple plasmids at different copy numbers requiring the use of several antibiotics. Mutations in the plasmid can destroy transcription of the antibiotic resistance genes and interject with resources management in the cell leading to necrosis. The selected colony is set on a glass-bottom Petri dish and a few focus planes are selected for microscopy tracking in both bright field and fluorescent domains. The protocol maintains the image focus for more than 12 hours under initial conditions that cannot be regulated, creating a few difficulties. For example, dead cells start to accumulate in the lenses' field of focus after a few hours of imaging, which causes toxins to buildup and the signal to blur and decay. Depletion of nutrients introduces new metabolic processes and hinder the desired response of the circuit. The experiment's temperature lowers the effectivity of inducers and antibiotics, which can further damage the reliability of the signal. The minimal media gel shrinks and dries, and as a result the optical focus changes over time. We developed this method to overcome these challenges in Escherichia coli, similar to previous works developing analogous methods for other micro-organisms. In addition, this method offers an algorithm to quantify the total stochastic noise in unaltered and altered cells, finding that the results are consistent with flow analyzer predictions as shown by a similar coefficient of variation (CV).

Continuous Measurement of Biological Noise in Escherichia Coli Using Time-lapse Microscopy

This work presents a microscopy method that allows live imaging of a single cell of Escherichia coli for analysis and quantification of the stochastic behavior of synthetic gene circuits.

The protocol developed here offers a tool to enable computer tracking of Escherichia coli division and fluorescent levels over several hours. The process ...

Functional Transcranial Doppler Ultrasound for Monitoring Cerebral Blood Flow

Functional transcranial Doppler ultrasound (fTCD) is the use of transcranial Doppler ultrasound (TCD) to study neural activation occurring during stimuli such as physical movement, activation of tactile sensors in the skin, and viewing images. Neural activation is inferred from an increase in the cerebral blood flow velocity (CBFV) supplying the region of the brain involved in processing sensory input. For example, viewing bright light causes increased neural activity in the occipital lobe of the cerebral cortex, leading to increased blood flow in the posterior cerebral artery, which supplies the occipital lobe. In fTCD, changes in CBFV are used to estimate changes in cerebral blood flow (CBF).
With its high temporal resolution measurement of blood flow velocities in the major cerebral arteries, fTCD complements other established functional imaging techniques. The goal of this Methods paper is to give step-by-step instructions for using fTCD to perform a functional imaging experiment. First, the basic steps for identifying the middle cerebral artery (MCA) and optimizing the signal will be described. Next, placement of a fixation device for holding the TCD probe in place during the experiment will be described. Finally, the breath-holding experiment, which is a specific example of a functional imaging experiment using fTCD, will be demonstrated.

Functional transcranial Doppler ultrasound complements other functional imaging modalities, with its high temporal resolution measurement of stimulus-induced changes in cerebral blood flow within the basal cerebral arteries. This Methods paper gives step-by-step instructions for using functional transcranial Doppler ultrasound to perform a functional imaging experiment.&#160;

Functional transcranial Doppler ultrasound (fTCD) is the use of transcranial Doppler ultrasound (TCD) to study neural activation occurring during stimuli ...

Electric and Magnetic Field Devices for Stimulation of Biological Tissues

Electric fields (EFs) and magnetic fields (MFs) have been widely used by tissue engineering to improve cell dynamics such as proliferation, migration, differentiation, morphology, and molecular synthesis. However, variables such stimuli strength and stimulation times need to be considered when stimulating either cells, tissues or scaffolds. Given that EFs and MFs vary according to cellular response, it remains unclear how to build devices that generate adequate biophysical stimuli to stimulate biological samples. In fact, there is a lack of evidence regarding the calculation and distribution when biophysical stimuli are applied. This protocol is focused on the design and manufacture of devices to generate EFs and MFs and implementation of a computational methodology to predict biophysical stimuli distribution inside and outside of biological samples. The EF device was composed of two parallel stainless-steel electrodes located at the top and bottom of biological cultures. Electrodes were connected to an oscillator to generate voltages (50, 100, 150 and 200 Vp-p) at 60 kHz. The MF device was composed of a coil, which was energized with a transformer to generate a current (1 A) and voltage (6 V) at 60 Hz. A polymethyl methacrylate support was built to locate the biological cultures in the middle of the coil. The computational simulation elucidated the homogeneous distribution of EFs and MFs inside and outside of biological tissues. This computational model is a promising tool that can modify parameters such as voltages, frequencies, tissue morphologies, well plate types, electrodes and coil size to estimate the EFs and MFs to achieve a cellular response.

This protocol describes the step-by-step process to build both electrical and magnetic stimulators used to stimulate biological tissues. The protocol includes a guideline to simulate computationally electric and magnetic fields and manufacture of stimulator devices.

Electric fields (EFs) and magnetic fields (MFs) have been widely used by tissue engineering to improve cell dynamics such as proliferation, migration, ...

Rapid, Enzymatic Methods for Amplification of Minimal, Linear Templates for Protein Prototyping using Cell-Free Systems

This protocol describes the design of a minimal DNA template and the steps for enzymatic amplification, enabling rapid prototyping of assayable proteins in less than 24 h using cell-free expression. After receiving DNA from a vendor, the gene fragment is PCR-amplified, cut, circularized, and cryo-banked. A small amount of the banked DNA is then diluted and amplified significantly (up to 106x) using isothermal rolling circle amplification (RCA). RCA can yield microgram quantities of the minimal expression template from picogram levels of starting material (mg levels if all starting synthetic fragment is used). In this work, a starting amount of 20 pg resulted in 4 &#181;g of the final product. The resulting RCA product (concatemer of the minimal template) can be added directly to a cell-free reaction with no purification steps. Due to this method being entirely PCR-based, it may enable future high-throughput screening efforts when coupled with automated liquid handling systems.

The study describes a protocol for creating large (&#181;g-mg) quantities of DNA for protein screening campaigns from synthetic gene fragments without cloning or using living cells. The minimal template is enzymatically digested and circularized and then amplified using isothermal rolling circle amplification. Cell-free expression reactions could be performed with the unpurified product.

This protocol describes the design of a minimal DNA template and the steps for enzymatic amplification, enabling rapid prototyping of assayable proteins ...