S.E. acknowledges the support from Duke Endowment Special Initiative Fund, Wells Fargo Fund, PhRMA Foundation, as well as Startup Funds from the University of North Carolina, Charlotte.

1. Synthesis of BSA-Au(III) Complex
<ol>
	<li>Dissolve 25 mg of BSA in 1 mL of high-performance liquid chromatography (HPLC) grade water in a 5 mL reaction vial. 
	NOTE: The solution should appear clear.</li>
	<li>Dissolve gold (III) chloride trihydrate (chloroauric acid) to a concentration of 5 mM in HPLC grade water. 
	NOTE: The solution should appear yellow. Chloroauric acid solution prepared at this concentration will result in a BSA to Au ratio of 1:13.
	<ol>
		<li>Alternatively, prepare a solution of chloroauric acid with a concentration of anywhere between 0.38 mM (BSA:Au = 1:1) to 20 mM (BSA:Au = 1 : 50) in HPLC grade water. 
		NOTE: Different ratios of BSA to gold will result in drastically different red fluorescence patterns of the excitation-emission map.</li>
	</ol>
	</li>
	<li>Place the reaction vial of BSA in a 37 &#176;C water bath and vigorously stir at 750 rpm using a magnetic stirrer.</li>
	<li>Immediately after the stirring begins, add 1 mL of chloroauric acid to the solution. The color of the solution should transform from clear to yellow.</li>
	<li>Stir the mixture for 2 min at 37 &#176;C and at 750 rpm using a magnetic stirrer.</li>
	<li>Into the reaction vial, add 100 &#956;L of 1 M NaOH to the solution to bring the pH to 12. 
	NOTE: Immediately after NaOH is added, the solution should darken slightly to a yellow-brown and then turn back to yellow.</li>
	<li>Continue to stir at 750 rpm for 2 h and at 37 &#176;C. The solution should slowly change from yellow to a dark yellow/brown color. This color change indicates the formation of the red fluorescing BSA-Au(III) complex.</li>
	<li>Allow the sample to sit at room temperature for 2 days, and the solution will continue to darken to an amber brown and the fluorescence intensity will increase.
	<ol>
		<li>Alternatively, let the sample continue to stir at 37 &#176;C for 12 more h as the color of the solution evolves to an amber brown.</li>
	</ol>
	</li>
</ol>
2. Synthesis of BSA-Cu(II)-Au(III)
<ol>
	<li>Dissolve 25 mg of BSA in 1 mL of HPLC grade water. The solution should appear clear.</li>
	<li>Dissolve copper (II) chloride dihydrate in HPLC grade water to a concentration of 5 mM. The solution should appear light blue.</li>
	<li>Add 1 mL of the aqueous BSA solution to a 5 mL reaction vial and place in a water bath at 37 &#176;C. Stir the mixture at 750 rpm.</li>
	<li>Immediately add 0.5 mL of the copper (II) chloride dihydrate solution to the reaction vial and mix for 2 min. The solution will remain light blue.</li>
	<li>Add 75 &#956;L of 1 M NaOH to bring the pH to 12 and allow to mix for 2 h. The solution will become purple.</li>
	<li>Dissolve chloroauric acid in HPLC grade water to a concentration of 5 mM.</li>
	<li>Add 0.5 mL of aqueous chloroauric acid to the reaction vial and adjust the pH back to 12 using 1 M NaOH.</li>
	<li>Stir the reaction mixture for 2 h. 
	NOTE: The solution should evolve to a brown color.</li>
</ol>
3. Synthesis of BSA-Ni(II)-Au(III)
<ol>
	<li>Dissolve 25 mg of BSA in 1 mL of HPLC grade water. The solution should appear clear.</li>
	<li>Dissolve nickel (II) chloride hexahydrate in HPLC grade water to a concentration of 5 mM. The solution should appear light green.</li>
	<li>Add 1 mL of the aqueous BSA solution to a 5 mL reaction vial and place in a water bath at 37 oC. Stir the mixture at 750 rpm.</li>
	<li>Immediately add 0.5 mL of the nickel (II) chloride hexahydrate solution to the reaction vial and mix for 2 min. 
	NOTE: The solution will remain light green.</li>
	<li>Add 75 &#956;L of 1 M NaOH to bring the pH to 12 and allow to mix for 2 h. 
	NOTE: The solution will become dark yellow.</li>
	<li>Dissolve chloroauric acid in HPLC grade water to a concentration of 5 mM.</li>
	<li>Add 0.5 mL of the aqueous chloroauric acid to the reaction vial and adjust the pH back to 12.</li>
	<li>Stir the reaction mixture for 2 h. 
	NOTE: The solution should evolve to a brown color.</li>
</ol>
4. Synthesis of [Cys34-capped-BSA]-Au(III)
<ol>
	<li>Dissolve 2 mg of N-ethylmaleimide (NEM) in 1 mL of phosphate buffered saline (PBS, pH 7.4).</li>
	<li>Dissolve 2 mg of BSA in 1 mL of PBS-NEM solution.</li>
	<li>Transfer the solution to a 5 mL reaction vial and stir at 20 &#176;C at 500 rpm for 1 h.</li>
	<li>Dialyze the solution using 12 kDa dialysis tubing in 500 mL of PBS, stirring at 50 rpm with a magnetic stirrer overnight to remove unreacted NEM.</li>
	<li>Dissolve chloroauric acid in PBS to a concentration of 0.4 mM. 
	NOTE: The solution will be a faint yellow.</li>
	<li>Transfer the reaction vial to a water bath at 37 &#176;C. Stir at 750 rpm.</li>
	<li>Immediately add 1 mL of chloroauric acid solution to the reaction vial and allow to mix for 2 min.</li>
	<li>Add 75 &#956;L of 1 M NaOH to the reaction vial to bring the pH to 12 and allow to mix for 2 h.</li>
</ol>
5. Synthesis of [all-thiol-capped-BSA]-Au(III)
<ol>
	<li>Prepare a solution of 2 M urea and 50 mM ammonium bicarbonate (NH4HCO3, pH 8.0) in HPLC grade water.</li>
	<li>Dissolve 3.3 mg of BSA in 1 mL of the above solution and transfer to a 5 mL reaction vial.</li>
	<li>Make a stock solution of 0.25 M tris(2-carboxyethyl)phosphine (TCEP) by dissolving 62.5 mg of TCEP in 1 mL of HPLC water.</li>
	<li>Add the stock solution of TCEP to the reaction vial until the final concentration of TCEP is 8 mM.</li>
	<li>Incubate the solution in a water bath for 1 h at 50 &#176;C. Stir at 500 rpm using a magnetic stirrer.</li>
	<li>Allow the solution to completely cool to room temperature.</li>
	<li>Prepare a stock solution of 100 mM NEM by dissolving 12.5 mg of NEM in 1 mL of HPLC grade water.</li>
	<li>Add the stock solution of NEM to the reaction vial until the final concentration of NEM is 16 mM.</li>
	<li>Allow the solution to combine for 2 h at 20 &#176;C. Stir at 500 rpm.</li>
	<li>Dialyze the solution using a 12 kDa dialysis tubing, stirring the solution at 50 rpm with a magnetic stirrer overnight in 500 mL of 50 mM NH4HCO3 to remove excess TCEP, NEM, and urea.</li>
	<li>Move the reaction vial to a water bath at 37 &#176;C and stir at 750 rpm.</li>
	<li>Dissolve chloroauric acid in HPLC grade water to a concentration of 0.66 mM.</li>
	<li>Immediately add 1 mL of chloroauric acid solution to the reaction vial and allow to mix for 2 min.</li>
	<li>Add 1 M NaOH until the pH of the solution is 12 and allow the solution to continue to mix for 2 h.</li>
</ol>

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The authors have nothing to disclose.

The BSA-Au(III) compounds prepared at pH 12 exhibit red fluorescence at an emission wavelength of &#955;em= 640 nm when excited with ultraviolet (UV) light &#955;ex= 365 nm (Figure 1A). The emergence of red fluorescence is a slow process and will take a few days at room temperature to increase to a maximum intensity. Running the reaction at 37 &#176;C will yield the optimum results, though higher temperature can be used to produce the red fluorescence faster. Irreversible degradation of the protein can occur at temperatures above 45 &#176;C23. The adjustment of pH so that BSA transforms into its aged (pH &#62; 10) conformation21 (&#34;A-form&#34;) is critical for red fluorescence; pH is finely adjusted from neutral to basic to determine the threshold of the occurrence of the red fluorescence (Figure 1B, C). For maximum red fluorescent intensity, the pH should be adjusted above 11. For red fluorescence, the pH can be adjusted beyond 11, although extremely basic (pH &#62; 13) conditions can denature BSA and cause red fluorescence to disappear.
Varying the stoichiometric ratio of BSA and Au can illustrate the binding of Au to BSA. The fluorescence spectra of the BSA-Au(III) compounds vary depending on the stoichiometric ratios of BSA to Au (Figure 2). As the BSA to gold ratio is adjusted to 1:26, a maximum in red fluorescence intensity is observed at &#955;ex= 500 nm. On the other hand, as the BSA:gold ratio is adjusted to 1:7, red fluorescence is observed primarily at &#955;ex= 365 nm. No red fluorescence can be detected at a ratio of BSA to Au less than 1:7 or above 1:52. The minimum number of gold cations required to produce the red fluorescence is less than 7 and more than 1, and the maximum number for the loss of the red fluorescence is greater than 52 (Figure 2B, C). Additionally, the reduction of all aforementioned samples will occur under excess sodium borohydride, elucidating that all samples still contain cationic Au(III). Furthermore, the addition of excess amounts of gold beyond 20 mM can cause the solution to become too acidic and denature the protein. If protein denaturation occurs due to high acidity, reduce the concentration of BSA and Au relatively to mediate this issue.
Competitive binding of Au and other metal cations to BSA can illustrate the binding sites in BSA. It is known that Cu(II) and Ni(II) both bind to the Asp fragment at the N-terminus of BSA24,25,26,27. Through the addition of Cu(II), a strong binder to the Asp-fragment, followed by the addition of Au(III), the absorbance spectra of BSA-Cu(II)-Au(III) and the absorbance spectra of BSA-Au(III) and BSA-Cu(II) are the same - indicating that gold and copper do not compete for the same binding site at the Asp-fragment (Figure 3C). Ni(II) binds weakly to the Asp-fragment and therefore Au(III) competes with Ni(II) as gold is added; it has been observed that the absorbance spectra of BSA-Ni(II) and BSA-Au(III) does not correlate with that of BSA-Ni(II)-Au(III) (Figure 3F). Through the above protocol, one can show how Au(III) binds to the known binding site of BSA. This technique also requires the adjustment of pH above 11 and the technique is modified by adding twice the concentration to BSA but at half volume, thus this should be performed at low BSA to Au ratios to maintain the protein conformation.
Modifying Cysteine residues in BSA can further elucidate the Au binding sites. Gold is known to have a high affinity for thiol28 and BSA possesses a surface accessible thiol on (Cys34)21. Through the blocking of this thiol, secondary binding sites can be elucidated. The blocking of this cysteine is carried out prior to the addition of Au(III) to the sample and shows an altered fluorescence pattern of BSA-Au(III), indicating a possible transfer pathway involved in the mechanism of the red fluorescence (Figure 4A). It is imperative to add the thiol blocking agent, in this case NEM, at a neutral pH. Cleaving all disulfide bonds and the subsequent capping of their free thiol groups reveals no red fluorescence (Figure 4B). These results indicate that a disulfide bond is required to produce a red fluorescent complex.
We have demonstrated in various protocols, through the use of spectroscopic and protein chemistry techniques, a method to analyze the BSA-Au(III) complexes. Protein chemistry techniques presented herein have not been widely used in the protein-based nano-materials research14. These techniques can be generally applicable and be valuable to understand the metal binding process and the possible binding sites in other, if not all, proteins such as trypsin, pepsin, lysozyme, and transferrin17,29. Proteins are dynamic, yet very precise &#34;nano-materials&#34;. Detailed understanding of the metal binding site could pave the way towards new protein-based materials with controlled optical properties with myriad of potential applications.

A BSA-Au compound exhibiting an ultraviolet (UV)-excitable red fluorescence, with remarkable stokes shift, has been originally synthesized by Xie et al.1. The unique and stable red fluorescence can find various applications in fields such as sensing2,3,4, imaging5,6,7, or nanomedicine8,9,10,11,12,13. This compound has been studied extensively by many researchers in the field of nano-science in recent years14,15,16. The BSA-Au compound has been interpreted as Au25 nanoclusters. The goal of the presented method is to examine this compound in detail and to understand the origin of the red fluorescence. By following the presented approach, the presence of multiple Au binding sites, and the origin of fluorescence, alternative to the single-site nucleation of Au25 nanoclusters, can be illustrated. The same approach can be employed to study how other proteins17,18,19 complexed with Au(III) can change their intrinsic fluorescent properties.
The synthesis of the red-fluorescent BSA-Au compound requires a narrow control of the molar ratios of BSA to Au (BSA:Au) to maximize the intensity of the fluorescence and the location of the peaks in the excitation-emission map (EEM)20. It can be shown that multiple binding sites exist for Au(III) to bind, including the Asparagine fragment (or Asp fragment, the first four amino acid residues at the N-terminus of BSA)21,22. The 34th amino acid of BSA (Cys-34) is also shown to coordinate Au(III) and to be involved in the mechanism of the red fluorescence([Cys34-capped-BSA]-Au(III))20. Upon cleaving all Cys-Cys disulfide bonds and capping all thiols, red fluorescence is not produced ([all-thiol-capped-BSA]-Au(III)). This indicates the necessity of Cys-Cys disulfide bonds as the Au(III) binding site to produce the red fluorescence.
Protein chemistry techniques have not been widely used to study the BSA-Au(III) complexes in the nano-science community. However, it would be valuable to employ these techniques to understand certain aspects of these complexes, as well as to gain detailed understanding of the Au(III) binding sites in BSA. This article is intended to show some of these techniques.

<table>
	<tbody>
		<tr>
			<td>Bovine Serum Albumin (BSA), 96%</td>
			<td>Sigma-Aldrich</td>
			<td>A5611</td>
			<td></td>
		</tr>
		<tr>
			<td>gold (III) chloride trihydrate, 99.9%</td>
			<td>Sigma-Aldrich</td>
			<td>520918</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper (II) chloride dihydrate, 99.999%</td>
			<td>Sigma-Aldrich</td>
			<td>459097</td>
			<td></td>
		</tr>
		<tr>
			<td>Nickel (II) chloride hexahydrate, 99.9%</td>
			<td>Sigma-Aldrich</td>
			<td>654507</td>
			<td></td>
		</tr>
		<tr>
			<td>N-Ethylmaleimide (NEM), &#62;99.0%</td>
			<td>Sigma-Aldrich</td>
			<td>4259</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris(2-carboxyethyl)phosphine hydrochloride (TCEP), &#62;98.0%</td>
			<td>Sigma-Aldrich</td>
			<td>C4706</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide, &#62;98.0%</td>
			<td>Sigma-Aldrich</td>
			<td>S8045</td>
			<td></td>
		</tr>
		<tr>
			<td>Urea, 99.5%</td>
			<td>Chem-Implex Int&#39;l</td>
			<td>30142</td>
			<td></td>
		</tr>
		<tr>
			<td>Phospate buffered saline (PBS)</td>
			<td>Corning</td>
			<td>MT21040CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium bicarbonate, 99.5%</td>
			<td>Sigma-Aldrich</td>
			<td>9830</td>
			<td></td>
		</tr>
	</tbody>
</table>

luminophore formation in various conformations of bovine serum albumin by binding of gold(iii)

The purpose of the presented protocols is to study the process of Au(III) binding to BSA, yielding conformation change-induced red fluorescence (&#955;em = 640 nm) of BSA-Au(III) complexes. The method adjusts the pH to show that the emergence of the red fluorescence is correlated with the pH-induced equilibrium transitions of the BSA conformations. Red fluorescent BSA-Au(III) complexes can only be formed with an adjustment of pH at or above 9.7, which corresponds to the &#34;A-form&#34; conformation of BSA. The protocol to adjust the BSA to Au molar ratio and to monitor the time-course of the process of Au(III) binding is described. The minimum number of Au(III) per BSA, to produce the red fluorescence, is less than seven. We describe the protocol in steps to illustrate the presence of multiple Au(III) binding sites in BSA. First, by adding copper (Cu(II)) or nickel (Ni(II)) cations followed by Au(III), this method reveals a binding site for Au(III) that is not the red fluorophore. Second, by modifying BSA by thiol capping agents, another nonfluorophore-forming Au(III) binding site is revealed. Third, changing the BSA conformation by cleaving and capping of the disulfide bonds, the possible Au(III) binding site(s) are illustrated. The protocol described, to control the BSA conformations and Au(III) binding, can be generally applied to study the interactions of other proteins and metal cations.

From the fluorescence of the BSA-Au(III) complex, it has been observed that the conversion of the intrinsic blue fluorescence of BSA (&#955;em = 400 nm) to red fluorescence (&#955;em = 640 nm) occurs at about pH 9.7 through an equilibrium transition (Figure 1). EEM of BSA-Au(III) at different BSA to Au molar ratios is shown in Figure 2, and this data shows how altering the molar ratios yields the same emission wavelength at different excitation wavelengths. Cu(II), Ni(II), and Au(III) competitively bind to a known site (Asp fragment) in BSA (Figure 3). The Cys34-capped BSA shows a change in EEM peak patterns upon Au(III) binding, and these results show how altering of specific binding sites alters fluorescence patterns. The all-thiol-capped BSA shows no red fluorescence and reveals Cys-Cys disulfide bonds as possible binding sites to produce the red fluorophore (Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58141/58141fig1.jpg" /> 
Figure 1. Fluorescence of BSA-Au(III) and the conformational induced change from blue to red. (A) The absorbance and fluorescence (&#955;ex = 365 nm) of BSA-Au(III). (B) Red fluorescence beings to emerge at around pH 9.7, at which the conformation of BSA changes. (C) Blue fluorescence decays as red fluorescence emerges. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58141/58141fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58141/58141fig2.jpg" /> 
Figure 2. Ratio-metric excitation-emission map (EEM) measurements of BSA-Au(III). EEM of BSA-Au(III) complex synthesized using the standard protocol while adjusting the ratio of BSA to gold. (A) BSA at pH 12, (B) BSA:Au = 1:1, (C) BSA:Au = 1:7, (D) BSA:Au = 1:13, (E) BSA:Au = 1:26, (F) BSA:Au = 1:30, (G) BSA:Au = 1:40, (H) BSA:Au = 1:52. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58141/58141fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58141/58141fig3.jpg" /> 
Figure 3. EEM of BSA-Cu(II)/Ni(II)-Au(III) complexes. Excitation-emission maps (excitation: 290 - 500 nm; emission: 300 - 850 nm) of BSA complexed with Cu(II)/Ni(II) at pH 12 (A and B), BSA complexed with Cu(II)/Ni(II) and then with Au(III) at pH 12 (C and D), and absorption spectra comparing BSA, BSA-Au, BSA-Cu(II)/Ni(II) and BSA-Cu(II)/Ni(II)-Au(III) (E and F). Curve 4 is compared with the superposition of Curves 2 and 3. This figure has been modified from Dixon, J. M. &#38; Egusa, S. J. Am. Chem. Soc. 140 2265-2271, (2018). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58141/58141fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58141/58141fig4.jpg" /> 
Figure 4. EEM of Cys34 capped and all thiols capped. EEM (excitation: 300 - 500 nm; emission: 300 - 700 nm) of (A) Cys34-capped BSA reacted with Au at pH 12. In (B), all Cys-Cys disulfide bonds in BSA were cleaved and then the all-thiol-capped-BSA was reacted with Au at pH 12. This figure has been modified from Dixon, J. M. &#38; Egusa, S. J. Am. Chem. Soc. 140 2265-2271, (2018). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58141/58141fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Luminophore Formation in Various Conformations of Bovine Serum Albumin by Binding of GoldIII at JoVE.com

Luminophore Formation in Various Conformations of Bovine Serum Albumin by Binding of GoldIII

The protocols for studying the binding of gold cations (Au(III)) to various conformations of bovine serum albumin (BSA) as well as for characterizing the conformational dependent unique BSA-Au fluorescence are presented.

Luminophore Formation in Various Conformations of Bovine Serum Albumin by Binding of Gold(III)

The purpose of the presented protocols is to study the process of Au(III) binding to BSA, yielding conformation change-induced red fluorescence (&#955;em ...

luminophore-formation-various-conformations-bovine-serum-albumin

Nanyang Technological University

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Chemistry

6.8K Views.  University of North Carolina. The protocols for studying the binding of gold cations (Au(III)) to various conformations of bovine serum albumin (BSA) as well as for characterizing the conformational dependent unique BSA-Au fluorescence are presented.

Video: Luminophore Formation in Various Conformations of Bovine Serum Albumin by Binding of GoldIII

LabVIEW-operated Novel Nanoliter Osmometer for Ice Binding Protein Investigations

Ice-binding proteins (IBPs), including antifreeze proteins, ice structuring proteins, thermal hysteresis proteins, and ice recrystallization inhibition proteins, are found in cold-adapted organisms and protect them from freeze injuries by interacting with ice crystals. IBPs are found in a variety of organism, including fish1, plants2, 3, arthropods4, 5, fungi6, and bacteria7. IBPs adsorb to the surfaces of ice crystals and prevent water molecules from joining the ice lattice at the IBP adsorption location. Ice that grows on the crystal surface between the adsorbed IBPs develops a high curvature that lowers the temperature at which the ice crystals grow, a phenomenon referred to as the Gibbs-Thomson effect. This depression creates a gap (thermal hysteresis, TH) between the melting point and the nonequilibrium freezing point, within which ice growth is arrested8-10, see Figure 1. One of the main tools used in IBP research is the nanoliter osmometer, which facilitates measurements of the TH activities of IBP solutions. Nanoliter osmometers, such as the Clifton instrument (Clifton Technical Physics, Hartford, NY,) and Otago instrument (Otago Osmometers, Dunedin, New Zealand), were designed to measure the osmolarity of a solution by measuring the melting point depression of droplets with nanoliter volumes. These devices were used to measure the osmolarities of biological samples, such as tears11, and were found to be useful in IBP research. Manual control over these nanoliter osmometers limited the experimental possibilities. Temperature rate changes could not be controlled reliably, the temperature range of the Clifton instrument was limited to 4,000 mOsmol (about -7.5 &deg;C), and temperature recordings as a function of time were not an available option for these instruments.
We designed a custom-made computer-controlled nanoliter osmometer system using a LabVIEW platform (National Instruments). The cold stage, described previously9, 10, contains a metal block through which water circulates, thereby functioning as a heat sink, see Figure 2. Attached to this block are thermoelectric coolers that may be driven using a commercial temperature controller that can be controlled via LabVIEW modules, see Figure 3. Further details are provided below. The major advantage of this system is its sensitive temperature control, see Figure 4. Automated temperature control permits the coordination of a fixed temperature ramp with a video microscopy output containing additional experimental details.
To study the time dependence of the TH activity, we tested a 58 kDa hyperactive IBP from the Antarctic bacterium Marinomonas primoryensis (MpIBP)12. This protein was tagged with enhanced green fluorescence proteins (eGFP) in a construct developed by Peter Davies' group (Queens University)10. We showed that the temperature change profile affected the TH activity. Excellent control over the temperature profile in these experiments significantly improved the TH measurements. The nanoliter osmometer additionally allowed us to test the recrystallization inhibition of IBPs5, 13. In general, recrystallization is a phenomenon in which large crystals grow larger at the expense of small crystals. IBPs efficiently inhibit recrystallization, even at low concentrations14, 15. We used our LabVIEW-controlled osmometer to quantitatively follow the recrystallization of ice and to enforce a constant ice fraction using simultaneous real-time video analysis of the images and temperature feedback from the sample chamber13. The real-time calculations offer additional control options during an experimental procedure. A stage for an inverted microscope was developed to accommodate temperature-controlled microfluidic devices, which will be described elsewhere16.
The Cold Stage System
The cold stage assembly (Figure 2) consists of a set of thermoelectric coolers that cool a copper plate. Heat is removed from the stage by flowing cold water through a closed compartment under the thermoelectric coolers. A 4 mm diameter hole in the middle of the copper plate serves as a viewing window. A 1 mm diameter in-plane hole was drilled to fit the thermistor. A custom-made copper disc (7 mm in diameter) with several holes (500 &mu;m in diameter) was placed on the copper plate and aligned with the viewing window. Air was pumped at a flow rate of 35 ml/sec and dried using Drierite (W.A. Hammond). The dry air was used to ensure a dry environment at the cooling stage. The stage was connected via a 9 pin connection outlet to a temperature controller (Model 3040 or 3150, Newport Corporation, Irvine, California, US). The temperature controller was connected via a cable to a computer GPIB-PCI card (National instruments, Austin, Texas, USA).

Ice binding proteins (IBPs), also known as antifreeze proteins, inhibit ice growth and are a promising additive for use in the cryopreservation of tissues. The main tool used to investigate IBPs is the nanoliter osmometer. We developed a home-designed cooling stage mounted on an optical microscope and controlled using a custom-built LabVIEW routine. The nanoliter osmometer described here manipulated the sample temperature in an ultra-sensitive manner.

Ice-binding proteins (IBPs), including antifreeze proteins, ice structuring proteins, thermal hysteresis proteins, and ice recrystallization inhibition ...

Large Scale Non-targeted Metabolomic Profiling of Serum by Ultra Performance Liquid Chromatography-Mass Spectrometry (UPLC-MS)

Non-targeted metabolite profiling by ultra performance liquid chromatography coupled with mass spectrometry (UPLC-MS) is a powerful technique to investigate metabolism. The approach offers an unbiased and in-depth analysis that can enable the development of diagnostic tests, novel therapies, and further our understanding of disease processes. The inherent chemical diversity of the metabolome creates significant analytical challenges and there is no single experimental approach that can detect all metabolites. Additionally, the biological variation in individual metabolism and the dependence of metabolism on environmental factors necessitates large sample numbers to achieve the appropriate statistical power required for meaningful biological interpretation. To address these challenges, this tutorial outlines an analytical workflow for large scale non-targeted metabolite profiling of serum by UPLC-MS. The procedure includes guidelines for sample organization and preparation, data acquisition, quality control, and metabolite identification and will enable reliable acquisition of data for large experiments and provide a starting point for laboratories new to non-targeted metabolite profiling by UPLC-MS.

Large Scale Non-targeted Metabolomic Profiling of Serum by Ultra Performance Liquid Chromatography-Mass Spectrometry UPLC-MS

Non-targeted metabolite profiling by ultra performance liquid chromatography coupled with mass spectrometry (UPLC-MS) is a powerful technique to investigate metabolism. This article outlines a typical workflow utilized for non-targeted metabolite profiling of serum including sample organization and preparation, data acquisition, data analysis, quality control, and metabolite identification.

Non-targeted metabolite profiling by ultra performance liquid chromatography coupled with mass spectrometry (UPLC-MS) is a powerful technique to ...

Covalent Binding of BMP-2 on Surfaces Using a Self-assembled Monolayer Approach

Bone morphogenetic protein 2 (BMP-2) is a growth factor embedded in the extracellular matrix of bone tissue. BMP-2 acts as trigger of mesenchymal cell differentiation into osteoblasts, thus stimulating healing and de novo bone formation. The clinical use of recombinant human BMP-2 (rhBMP-2) in conjunction with scaffolds has raised recent controversies, based on the mode of presentation and the amount to be delivered. The protocol presented here provides a simple and efficient way to deliver BMP-2 for in vitro studies on cells. We describe how to form a self-assembled monolayer consisting of a heterobifunctional linker, and show the subsequent binding step to obtain covalent immobilization of rhBMP-2. With this approach it is possible to achieve a sustained presentation of BMP-2 while maintaining the biological activity of the protein. In fact, the surface immobilization of BMP-2 allows targeted investigations by preventing unspecific adsorption, while reducing the amount of growth factor and, most notably, hindering uncontrolled release from the surface. Both short- and long-term signaling events triggered by BMP-2 are taking place when cells are exposed to surfaces presenting covalently immobilized rhBMP-2, making this approach suitable for in vitro studies on cell responses to BMP-2 stimulation.

We describe a method for accomplishing efficient immobilization of BMP-2 on surfaces. Our approach is based on the formation of a self-assembled monolayer to achieve the covalent binding of BMP-2 via its free amine residues. This method is a useful tool to study signaling at the cell membrane.

Bone morphogenetic protein 2 (BMP-2) is a growth factor embedded in the extracellular matrix of bone tissue. BMP-2 acts as trigger of mesenchymal cell ...

Microwave-assisted Functionalization of Poly(ethylene glycol) and On-resin Peptides for Use in Chain Polymerizations and Hydrogel Formation

One of the main benefits to using poly(ethylene glycol) (PEG) macromers in hydrogel formation is synthetic versatility. The ability to draw from a large variety of PEG molecular weights and configurations (arm number, arm length, and branching pattern) affords researchers tight control over resulting hydrogel structures and properties, including Young&#8217;s modulus and mesh size. This video will illustrate a rapid, efficient, solvent-free, microwave-assisted method to methacrylate PEG precursors into poly(ethylene glycol) dimethacrylate (PEGDM). This synthetic method provides much-needed starting materials for applications in drug delivery and regenerative medicine. The demonstrated method is superior to traditional methacrylation methods as it is significantly faster and simpler, as well as more economical and environmentally friendly, using smaller amounts of reagents and solvents. We will also demonstrate an adaptation of this technique for on-resin methacrylamide functionalization of peptides. This on-resin method allows the N-terminus of peptides to be functionalized with methacrylamide groups prior to deprotection and cleavage from resin. This allows for selective addition of methacrylamide groups to the N-termini of the peptides while amino acids with reactive side groups (e.g.&#160;primary amine of lysine, primary alcohol of serine, secondary alcohols of threonine, and phenol of tyrosine) remain protected, preventing functionalization at multiple sites. This article will detail common analytical methods (proton Nuclear Magnetic Resonance spectroscopy (;H-NMR) and Matrix Assisted Laser Desorption Ionization Time of Flight mass spectrometry (MALDI-ToF)) to assess the efficiency of the functionalizations. Common pitfalls and suggested troubleshooting methods will be addressed, as will modifications of the technique which can be used to further tune macromer functionality and resulting hydrogel physical and chemical properties. Use of synthesized products for the formation of hydrogels for drug delivery and cell-material interaction studies will be demonstrated, with particular attention paid to modifying hydrogel composition to affect mesh size, controlling hydrogel stiffness and drug release.

Microwave-assisted Functionalization of Polyethylene glycol and On-resin Peptides for Use in Chain Polymerizations and Hydrogel Formation

This video will illustrate a rapid, efficient method to methacrylate poly(ethylene glycol), enabling chain polymerizations and hydrogel synthesis. It will demonstrate how to similarly introduce methacrylamide functionalities into peptides, detail common analytical methods to assess functionalization efficiency, provide suggestions for troubleshooting and advanced modifications, and demonstrate typical hydrogel characterization techniques.

One of the main benefits to using poly(ethylene glycol) (PEG) macromers in hydrogel formation is synthetic versatility. The ability to draw from a large ...

Formation of Ordered Biomolecular Structures by the Self-assembly of Short Peptides

In nature, complex functional structures are formed by the self-assembly of biomolecules under mild conditions. Understanding the forces that control self-assembly and mimicking this process in vitro will bring about major advances in the areas of materials science and nanotechnology. Among the available biological building blocks, peptides have several advantages as they present substantial diversity, their synthesis in large scale is straightforward, and they can easily be modified with biological and chemical entities1,2. Several classes of designed peptides such as cyclic peptides, amphiphile peptides and peptide-conjugates self-assemble into ordered structures in solution. Homoaromatic dipeptides, are a class of short self-assembled peptides that contain all the molecular information needed to form ordered structures such as nanotubes, spheres and fibrils3-8. A large variety of these peptides is commercially available.
This paper presents a procedure that leads to the formation of ordered structures by the self-assembly of homoaromatic peptides. The protocol requires only commercial reagents and basic laboratory equipment. In addition, the paper describes some of the methods available for the characterization of peptide-based assemblies. These methods include electron and atomic force microscopy and Fourier-Transform Infrared Spectroscopy (FT-IR). Moreover, the manuscript demonstrates the blending of peptides (coassembly) and the formation of a &#34;beads on a string&#34;-like structure by this process.9 The protocols presented here can be adapted to other classes of peptides or biological building blocks and can potentially lead to the discovery of new peptide-based structures and to better control of their assembly.

This paper describes the formation of highly ordered peptide-based structures by the spontaneous process of self-assembly. The method utilizes commercially available peptides and common lab equipment. This technique can be applied to a large variety of peptides and may lead to the discovery of new peptide-based assemblies.

In nature, complex functional structures are formed by the self-assembly of biomolecules under mild conditions. Understanding the forces that control ...

Determining the Ice-binding Planes of Antifreeze Proteins by Fluorescence-based Ice Plane Affinity

Antifreeze proteins (AFPs) are expressed in a variety of cold-hardy organisms to prevent or slow internal ice growth. AFPs bind to specific planes of ice through their ice-binding surfaces. Fluorescence-based ice plane affinity (FIPA) analysis is a modified technique used to determine the ice planes to which the AFPs bind. FIPA is based on the original ice-etching method for determining AFP-bound ice-planes. It produces clearer images in a shortened experimental time. In FIPA analysis, AFPs are fluorescently labeled with a chimeric tag or a covalent dye then slowly incorporated into a macroscopic single ice crystal, which has been preformed into a hemisphere and oriented to determine the a- and c-axes. The AFP-bound ice hemisphere is imaged under UV light to visualize AFP-bound planes using filters to block out nonspecific light. Fluorescent labeling of the AFPs allows real-time monitoring of AFP adsorption into ice. The labels have been found not to influence the planes to which AFPs bind. FIPA analysis also introduces the option to bind more than one differently tagged AFP on the same single ice crystal to help differentiate their binding planes. These applications of FIPA are helping to advance our understanding of how AFPs bind to ice to halt its growth and why many AFP-producing organisms express multiple AFP isoforms.

Antifreeze proteins (AFPs) bind to specific planes of ice to prevent or slow ice growth. Fluorescence-based ice plane affinity (FIPA) analysis is a modification of the original ice-etching method for determination of AFP-bound ice planes. AFPs are fluorescently labeled, incorporated into macroscopic single ice crystals, and visualized under UV light.

Antifreeze proteins (AFPs) are expressed in a variety of cold-hardy organisms to prevent or slow internal ice growth. AFPs bind to specific planes of ice ...

Water in Oil Emulsions: A New System for Assembling Water-soluble Chlorophyll-binding Proteins with Hydrophobic Pigments

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. Their functionality critically depends on their specific organization within large and elaborate multisubunit transmembrane protein complexes. In order to understand at the molecular level how these complexes facilitate solar energy conversion, it is essential to understand protein-pigment, and pigment-pigment interactions, and their effect on excited dynamics. One way of gaining such understanding is by constructing and studying complexes of Chls with simple water-soluble recombinant proteins. However, incorporating the lipophilic Chls and BChls into water-soluble proteins is difficult. Moreover, there is no general method, which could be used for assembly of water-soluble proteins with hydrophobic pigments. Here, we demonstrate a simple and high throughput system based on water-in-oil emulsions, which enables assembly of water-soluble proteins with hydrophobic Chls. The new method was validated by assembling recombinant versions of the water-soluble chlorophyll binding protein of Brassicaceae plants (WSCP) with Chl a. We demonstrate the successful assembly of Chl a using crude lysates of WSCP expressing E. coli cell, which may be used for developing a genetic screen system for novel water-soluble Chl-binding proteins, and for studies of Chl-protein interactions and assembly processes.

This manuscript describes a simple and high-throughput method for assembling water-soluble proteins with hydrophobic pigments that is based on water-in-oil emulsions. We demonstrate the effectiveness of the method in the assembly of native chlorophylls with four variants of recombinant water-soluble-chlorophyll binding proteins (WSCPs) of Brassica plants expressed in E. coli.

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. ...

Radiolabeling and Quantification of Cellular Levels of Phosphoinositides by High Performance Liquid Chromatography-coupled Flow Scintillation

Phosphoinositides (PtdInsPs) are essential signaling lipids responsible for recruiting specific effectors and conferring organelles with molecular identity and function. Each of the seven PtdInsPs varies in their distribution and abundance, which are tightly regulated by specific kinases and phosphatases. The abundance of PtdInsPs can change abruptly in response to various signaling events or disturbance of the regulatory machinery. To understand how these events lead to changes in the amount of PtdInsPs and their resulting impact, it is important to quantify PtdInsP levels before and after a signaling event or between control and abnormal conditions. However, due to their low abundance and similarity, quantifying the relative amounts of each PtdInsP can be challenging. This article describes a method for quantifying PtdInsP levels by metabolically labeling cells with 3H-myo-inositol, which is incorporated into PtdInsPs. Phospholipids are then precipitated and deacylated. The resulting soluble 3H-glycero-inositides are further extracted, separated by high-performance liquid chromatography (HPLC), and detected by flow scintillation. The labeling and processing of yeast samples is described in detail, as well as the instrumental setup for the HPLC and flow scintillator. Despite losing structural information regarding acyl chain content, this method is sensitive and can be optimized to concurrently quantify all seven PtdInsPs in cells.

Phosphoinositides are signaling lipids whose relative abundance rapidly changes in response to various stimuli. This article describes a method to measure the abundance of phosphoinositides by metabolically labeling cells with 3H-myo-inositol, followed by extraction and deacylation. Extracted glycero-inositides are then separated by high-performance liquid chromatography and quantified by flow scintillation.

Phosphoinositides (PtdInsPs) are essential signaling lipids responsible for recruiting specific effectors and conferring organelles with molecular ...

In Situ Characterization of Hydrated Proteins in Water by SALVI and ToF-SIMS

This work demonstrates in situ characterization of protein biomolecules in the aqueous solution using the System for Analysis at the Liquid Vacuum Interface (SALVI) and time-of-flight secondary ion mass spectrometry (ToF-SIMS). The fibronectin protein film was immobilized on the silicon nitride (SiN) membrane that forms the SALVI detection area. During ToF-SIMS analysis, three modes of analysis were conducted including high spatial resolution mass spectrometry, two-dimensional (2D) imaging, and depth profiling. Mass spectra were acquired in both positive and negative modes. Deionized water was also analyzed as a reference sample. Our results show that the fibronectin film in water has more distinct and stronger water cluster peaks compared to water alone. Characteristic peaks of amino acid fragments are also observable in the hydrated protein ToF-SIMS spectra. These results illustrate that protein molecule adsorption on a surface can be studied dynamically using SALVI and ToF-SIMS in the liquid environment for the first time.

In Situ Characterization of Hydrated Proteins in Water by SALVI and ToF-SIMS

This work presents a protocol for liquid handling and sample introduction to a microchannel for in situ time-of-flight secondary ion mass spectrometry analysis of protein biomolecules in an aqueous solution.

This work demonstrates in situ characterization of protein biomolecules in the aqueous solution using the System for Analysis at the Liquid Vacuum ...

DNA-magnetic Particle Binding Analysis by Dynamic and Electrophoretic Light Scattering

Isolation of DNA using magnetic particles is a field of high importance in biotechnology and molecular biology research. This protocol describes the evaluation of DNA-magnetic particles binding via dynamic light scattering (DLS) and electrophoretic light scattering (ELS). Analysis by DLS provides valuable information on the physicochemical properties of particles including particle size, polydispersity, and zeta potential. The latter describes the surface charge of the particle which plays major role in electrostatic binding of materials such as DNA. Here, a comparative analysis exploits three chemical modifications of nanoparticles and microparticles and their effects on DNA binding and elution. Chemical modifications by branched polyethylenimine, tetraethyl orthosilicate and (3-aminopropyl)triethoxysilane are investigated. Since DNA exhibits a negative charge, it is expected that zeta potential of particle surface will decrease upon binding of DNA. Forming of clusters should also affect particle size. In order to investigate the efficiency of these particles in isolation and elution of DNA, the particles are mixed with DNA in low pH (~6), high ionic strength and dehydration environment. Particles are washed on magnet and then DNA is eluted by Tris-HCl buffer (pH = 8). DNA copy number is estimated using quantitative polymerase chain reaction (PCR). Zeta potential, particle size, polydispersity and quantitative PCR data are evaluated and compared. DLS is an insightful and supporting method of analysis that adds a new perspective to the process of screening of particles for DNA isolation.

This protocol describes the synthesis of magnetic particles and evaluation of their DNA-binding properties via dynamic and electrophoretic light scattering. This method focuses on monitoring changes in particle size, their polydispersity and zeta potential of particle surface which play major role in binding of materials such as DNA.

Isolation of DNA using magnetic particles is a field of high importance in biotechnology and molecular biology research. This protocol describes the ...

Luminophore Formation in Various Conformations of Bovine Serum Albumin by Binding of Gold(III)

Summary

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Chapters in this video

LabVIEW-operated Novel Nanoliter Osmometer for Ice Binding Protein Investigations

Large Scale Non-targeted Metabolomic Profiling of Serum by Ultra Performance Liquid Chromatography-Mass Spectrometry (UPLC-MS)

Covalent Binding of BMP-2 on Surfaces Using a Self-assembled Monolayer Approach

Microwave-assisted Functionalization of Poly(ethylene glycol) and On-resin Peptides for Use in Chain Polymerizations and Hydrogel Formation

Formation of Ordered Biomolecular Structures by the Self-assembly of Short Peptides

Determining the Ice-binding Planes of Antifreeze Proteins by Fluorescence-based Ice Plane Affinity

Water in Oil Emulsions: A New System for Assembling Water-soluble Chlorophyll-binding Proteins with Hydrophobic Pigments

Radiolabeling and Quantification of Cellular Levels of Phosphoinositides by High Performance Liquid Chromatography-coupled Flow Scintillation

In Situ Characterization of Hydrated Proteins in Water by SALVI and ToF-SIMS

DNA-magnetic Particle Binding Analysis by Dynamic and Electrophoretic Light Scattering