This work was supported by National Science Foundation through NSF OIA-1833040. The National High Magnetic Field Laboratory (NHMFL) is supported by National Science Foundation through DMR-1157490 and the State of Florida. The MAS-DNP system at NHMFL is funded in part by NIH S10 OD018519 and NSF CHE-1229170.

1. Growth of 13C, 15N-labeled Aspergillus fumigatus Liquid Medium
<ol>
	<li>Preparation of unlabeled and 13C, 15N-labeled growth medium 
	NOTE: Both Yeast Extract Peptone Dextrose medium (YPD) and the improved minimal medium43 were used for the maintenance of fungal culture. All steps after autoclaving are performed in a laminar flow hood to minimize contamination.
	<ol>
		<li>Preparation of unlabeled liquid medium: Dissolve 6.5 g of YPD powder in 100 mL of distilled water and then autoclave for 25 min at 134 &#176;C.</li>
		<li>Preparation of unlabeled solid medium
		<ol>
			<li>Add 1.5 g of agar and 6.5 g of YPD powder in 100 mL of distilled water.</li>
			<li>Autoclave the medium for 25 min at 121 &#176;C and then cool down to approximately 50 &#176;C.</li>
			<li>Transfer 13-15 mL of the medium into each pre-sterile plastic Petri dish and cover the dish using a lid immediately.</li>
		</ol>
		</li>
		<li>Preparation of 13C, 15N-labeled liquid medium 
		NOTE: To prepare the growth solution for isotope labeling, a minimal medium containing 13C-glucose and 15N-sodium nitrate and a trace-element solution are prepared separately and then mixed before use.
		<ol>
			<li>Prepare 100 mL of the isotope-containing minimal medium as listed in Table 1. Adjust the pH to 6.6 using NaOH (1 M) or HCl (1M) solution.</li>
			<li>Autoclave the minimal medium for 25 min at 121&#160;&#176;C.</li>
			<li>Prepare 100 mL (1,000x) of trace elements solution, dissolve the salts listed in Table 2 in the distilled water. Autoclave the solution for 25 min at 134 &#176;C. Cool down and store the solution at 4 &#176;C for short-term use. The pH will be about 6.5 and can be checked using a pH meter.</li>
			<li>Add 0.1 mL of trace elements solution to 100 mL of 13C, 15N-labeled minimal medium as listed in Table 2 before use.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Growth of the fungal materials
	<ol>
		<li>Transfer a small amount of fungi from the storage onto a YPD plate using an inoculating loop in a laminar flow hood. Keep the culture at 30 &#176;C for 2 days in an incubator.</li>
		<li>Use an inoculating loop to transfer an active growing fungal edge to the 13C,15N-labeling solution in a laminar flow hood. Keep the culture at 30 &#176;C for 3-5 days at 220 rpm in a shaking incubator.</li>
		<li>Centrifuge at 4,000 x g for 20 min. Remove the supernatant and collect the pellet.</li>
		<li>Use a tweezer to collect ~0.5 g of well-hydrated pellet (&#62;50 wt% hydration) for NMR studies. Loss of hydration at any point will substantially worsen the spectral resolution. 
		NOTE: If needed, a small amount (0.1 g) of the hydrated mycelia can be separated and fully dried under N2 gas flow in a hood or a lyophilizer to estimate the hydration level and calculate the dry mass percentage. Usually, a pellet containing ~0.3 g dry mass can be obtained after 3 days. If the NMR experiment to be conducted is long (&#62;7 days) and/or if the state of the fungi needs to be fixed, the fungal material can be deeply frozen in liquid N2 for 10-20 min before further processing. If the experiment will be short (3-6 days), the freezing can be skipped so that the sample can remain fresh.</li>
		<li>Mix the excess material with 20% (v/v) of glycerol in a centrifuge tube and keep it in a -80 &#7506;C freezer for long-term storage.</li>
	</ol>
	</li>
</ol>
2. Preparation of A. fumigatus for Solid-state NMR and DNP Studies
<ol>
	<li>Preparation of A. fumigatus for solid-state NMR experiments
	<ol>
		<li>Dialyze the 13C, 15N-labeled fungal sample (Step 1.2.4.) against 1 L of 10 mM phosphate buffer (pH 7.0) at 4&#176;C using a dialysis bag with a 3.5 kDa molecular weight cutoff to remove small molecules from the growth medium for a total period of 3 days. Change the buffer twice daily. 
		NOTE: Alternatively, the sample could be washed for 6-10 times using deionized water to remove residual small molecules.</li>
		<li>Transfer the sample into a 15 mL tube and centrifuge for 5 min (10,000 x g) using a benchtop centrifuge. Remove the supernatant and collect the remaining fungal materials.</li>
		<li>Pack 70-80 mg of the uniformly 13C-labeled and well-hydrated sample paste into a 4-mm ZrO2 rotor or 30-50 mg to 3.2 mm rotors for NMR experiments. Repetitively squeeze the sample gently using a metal rod and absorb the excess water using paper.</li>
		<li>Tightly cap the rotor and insert the sample into the spectrometer for solid-state NMR characterization. 
		NOTE: The brand-new rotors are suggested to minimize the possibility of rotor crash and sample spill in the NMR spectrometer. If needed, a disposable Kel-F insert with sealing screws can be used to serve as a secondary container inside the rotor.</li>
	</ol>
	</li>
	<li>Preparation of A. fumigatus samples for DNP experiments
	<ol>
		<li>Prepare 100 &#181;L of DNP solvents29,44 (also known as the DNP matrix) in a 1.5 mL microcentrifuge tube for 13C,15N-labeled fungal samples. This DNP matrix contains a mixture of d8-glycerol/D2O/H2O (60/30/10 Vol%). 
		NOTE: If unlabeled samples are to be investigated, then prepare the DNP matrix using 13C-depleted d8-glycerol (12C3, 99.95%; D8, 98%) and D2O and H2O to avoid 13C signal contribution from the solvents.</li>
		<li>Dissolve 0.7 mg of AMUPol45 in 100 &#181;L of DNP solvents to form 10 mM radical stock solution. Vortex for 2-3 min to ensure that radicals are fully dissolved in the solution.</li>
		<li>Soak 10 mg of the dialyzed 13C, 15N-labeled fungal materials as described in prior steps (Steps 2.1.1 and 2.1.2) into 50 &#181;L of AMUPol solution, and mildly grind the mixture using a pestle and a mortar to ensure penetration of the radicals into the porous cell walls. 
		NOTE: To reduce the rate of hydration loss, the grinding can also take place in a microcentrifuge tube using a micropestle.</li>
		<li>Add another 30 &#181;L of the radical solution to the grinded pellet to further hydrate the fungal sample.</li>
		<li>Pack the pellet into a 3.2-mm sapphire rotor, squeeze mildly and remove the excess DNP solvent. Add a 3.2-mm silicone plug to prevent the loss of hydration. Typically, 5-30 mg of sample can be packed to the rotor. The exact amount needs to be determined by the sensitivity requirement of the NMR experiments to be conducted.</li>
		<li>Insert and spin up the sample in a DNP spectrometer, measure a DNP-enhanced spectrum under microwave irradiation and compare it with the microwave-off spectrum. This will lead to an enhancement factor &#949;on/off, which should be 20-40 for these complex materials. Run the designed experiments to determine cell wall structure.</li>
	</ol>
	</li>
</ol>
3. Preparation of Plant Biomass for NMR and DNP Studies
<ol>
	<li>Preparation of plant materials for solid-state NMR
	<ol>
		<li>Produce uniformly 13C-labeled plants in-house using 13CO2 supplies in a growth chamber or 13C-glucose medium as described previously46,47 or directly purchase labeled materials from isotope-labeling companies. 
		NOTE: 13C-glucose can only be used in dark growth to avoid the introduction of 12C by photosynthesis.</li>
		<li>Cut the uniformly 13C labeled plant material into small pieces (typically 1-2 mm in dimension) using a laboratory razor blade. 
		NOTE: Depending on the purpose, the extracted cell walls are sometimes used for structural characterization and the detailed protocols are reported in previous studies21,46.</li>
		<li>If the sample was previously dried, add 100 &#181;L of water to 30 mg of plant materials in a 1.5 mL microcentrifuge tube, vortex, equilibrate at room temperature for 1 day. Centrifuge at 4,000 x g for 10 min and remove the excess water using a pipette.</li>
		<li>If the sample was never-dried at any point, directly use the sample without further treatment.</li>
		<li>Pack the resulting plant materials into 3.2-mm or 4-mm ZrO2 rotors for solid-state NMR experiments.</li>
	</ol>
	</li>
	<li>Preparation of plant materials for DNP studies
	<ol>
		<li>Prepare 60 &#181;L stock solution of 10 mM AMUPol radical as described steps 2.2.1 and 2.2.2.</li>
		<li>Cut the uniformly 13C labeled plant material to be studied into small pieces (1-2 mm in dimension) using a laboratory razor blade and weigh 20 mg of the plant materials.</li>
		<li>Hand-grind the plant pieces into small particles (~1-2 mm in size) using a mortar and pestle. The final powders have a homogenous appearance.</li>
		<li>Add 40 &#181;L of the DNP stock solution prepared in prior step (Step 2.2.2) to the plant material and grind mildly for 5 min to ensure homogeneous mixing with the radical.</li>
		<li>Add another 20 &#181;L of the stock solution to further hydrate the plant material after grinding.</li>
		<li>Pack the equilibrated plant sample into a 3.2-mm sapphire rotor for DNP experiments. Insert a silicone plug to avoid the loss of hydration.</li>
	</ol>
	</li>
</ol>
4. Standard Solid-State NMR Experiments for Initial Characterization of Carbohydrate-Rich Biomaterials
NOTE: A brief overview of the NMR experiments is provided in this section. However, structural elucidation typically requires extensive expertise. Therefore, collaborative efforts with NMR spectroscopists is recommended.
<ol>
	<li>Measure 1D 13C Cross Polarization (CP), 13C Direct Polarization (DP) with 2-s and 35-s recycle delays, and 1H-13C INEPT48,49 spectra to obtain a general understanding of the dynamical distribution of cell components (Figure 1a). The cell walls are typically the relatively rigid portion and exhibit dominant signals in the CP spectrum.</li>
	<li>Measure a series of standard 2D 13C-13C correlation experiments for resonance assignments of 13C signals. Start with refocused INADEQUATE50,51 to obtain carbon connectivity, which need to be assisted by a series of through-space experiments such as 1.5-ms RFDR52 (Figure 1b) and 50-ms CORD/DARR53 experiments. 
	NOTE: If it is of interest to find a sample rich in a specific component, for example, the primary or secondary cell walls, then multiple segments or multiple plants may need to be measured separately to find the sample with the optimal composition.</li>
	<li>Conduct 2D 15N-13C correlation experiments which can be measured to facilitate the resonance assignments of proteins and nitrogenated carbohydrates. 
	NOTE: The resonance assignment is typically time-consuming. A method is currently being developed to facilitate the resonance assignment of carbohydrate signals for those scientists without prior experience.</li>
	<li>Measure more specialized experiments to determine the spatial proximities (Figure 1c, d), hydration and mobilities of complex biomolecules to determine the three-dimensional structure of the carbohydrate-rich materials as systematically described previously22,29.</li>
</ol>

We have nothing to disclose.

Compared with the biochemical methods, solid-state NMR has advantages as a non-destructive and high-resolution technique. NMR is also quantitative in compositional analysis, and unlike most other analytical methods, does not have the uncertainties introduced by the limited solubility of biopolymers. Establishment of the current protocol facilitates future studies on carbohydrate-rich biomaterials and functionalized polymers. However, it should be noted that the resonance assignment and data analysis can be time-consuming...

Carbohydrates play a central role in various biological processes such as energy storage, structural building, and cellular recognition and adhesion. They are enriched in the cell wall, which is a fundamental component in plants, fungi, algae and bacteria1,2,3. The cell wall serves as a central source for the production of biofuel and biomaterials, as well as a promising target for antimicrobial therapies4,5,6,7,8,9.
The contemporary understanding of these complex materials has been substantially advanced by decades of efforts that were devoted to the structural characterization using four major biochemical or genetic methods. The first major method relies on sequential treatments using harsh chemicals or enzymes to break down the cell walls into different portions, which is followed by compositional and linkage analysis of sugars in each fraction10. This method sheds light on the domain distribution of polymers, but the interpretation may be misleading due to the chemical and physical properties of biomolecules. For example, it is difficult to determine whether the alkali-extractable fraction originates from a single domain of less structured molecules or from spatially separated molecules with comparable solubility. Second, the extracted portions or whole cell walls can also be measured using solution NMR to determine the covalent linkages, also termed as crosslinking, between different molecules11,12,13,14,15. In this way, the detailed structure of covalent anchors could be probed, but limitations may exist due to the low solubility of polysaccharides, the relatively small number of crosslinking sites, and the ignorance of non-covalent effects that stabilizes polysaccharide packing, including the hydrogen-bonding, van der Waals force, electrostatic interaction and polymer entanglement. Third, the binding affinity has been determined in vitro using isolated polysaccharides16,17,18,19, but the purification procedures may substantially alter the structure and properties of these biomolecules. This method also fails to replicate the sophisticated deposition and assembly of macromolecules after biosynthesis. Finally, the phenotype, cell morphology and mechanical properties of genetic mutants with attenuated production of certain cell wall component shed lights on the structural functions of polysaccharides, but more molecular evidence is needed to bridge these macroscopic observations with the engineered function of protein machineries20.
Recent advances in the development and application of multidimensional solid-state NMR spectroscopy have introduced a unique opportunity for solving these structural puzzles. 2D/3D solid-state NMR experiments enable high-resolution investigation of the composition and architecture of carbohydrate-rich materials in the native state without major perturbation. Structural studies have been successfully conducted on both primary and secondary cell walls of plants, the catalytically treated biomass, bacterial biofilm, the pigment ghosts in fungi and, recently by the authors, the intact cell walls in a pathogenic fungus Aspergillus fumigatus21,22,23,24,25,26,27,28,29,30,31. The development of dynamic nuclear polarization (DNP)32,33,34,35,36,37,38,39,40,41,42 substantially facilitates NMR structural elucidation as the sensitivity enhancement by DNP markedly shortens the experimental time on these complex biomaterials. The protocol described here details the procedures for isotope-labeling the fungus A. fumigatus and preparing fungal and plant samples for solid-state NMR and DNP characterization. Similar labeling procedures should be applicable to other fungi with altered medium, and the sample preparation procedures should be generally applicable to other carbohydrate-rich biomaterials.

<table>
	<tbody>
		<tr>
			<td>Ammonium Molybdate Tetrahydrate</td>
			<td>Acros Organics</td>
			<td>12054-85-2</td>
			<td></td>
		</tr>
		<tr>
			<td>AMUPol</td>
			<td>Cortecnet</td>
			<td>C010P002</td>
			<td></td>
		</tr>
		<tr>
			<td>Analytical weighing balance</td>
			<td>Ohaus</td>
			<td>B730439218</td>
			<td>Model PA84C</td>
		</tr>
		<tr>
			<td>Bioclave 16 L</td>
			<td>VWR</td>
			<td>470230-598</td>
			<td></td>
		</tr>
		<tr>
			<td>Biosafety Cabinet</td>
			<td>Labconco corporation</td>
			<td>302319100</td>
			<td></td>
		</tr>
		<tr>
			<td>Boric acid</td>
			<td>VWR</td>
			<td>BDH9222</td>
			<td>store at 15-30 &#176;C</td>
		</tr>
		<tr>
			<td>Cobalt(II) Chloride Hexahydrate</td>
			<td>Honeywell|Fluka</td>
			<td>60820</td>
			<td>&#8805;98 %</td>
		</tr>
		<tr>
			<td>Copper(II) Sulfate Pentahydrate</td>
			<td>BDH</td>
			<td>BDH9312</td>
			<td>&#8805;98 %</td>
		</tr>
		<tr>
			<td>Corning LSE shaking incubator</td>
			<td>Thermo Fisher Scientific</td>
			<td>7202152</td>
			<td></td>
		</tr>
		<tr>
			<td>D2O</td>
			<td>Sigma Aldrich</td>
			<td>151882</td>
			<td>99.9 atom % D</td>
		</tr>
		<tr>
			<td>d6-DMSO</td>
			<td>Sigma Aldrich</td>
			<td>151874</td>
			<td>99.9 atom % D</td>
		</tr>
		<tr>
			<td>d8-glycerol</td>
			<td>Sigma Aldrich</td>
			<td>447498</td>
			<td>&#8805;99 atom % D</td>
		</tr>
		<tr>
			<td>Dialysis tubing 3.2 kDa</td>
			<td>Sigma Aldrich</td>
			<td>D2272</td>
			<td>132724</td>
		</tr>
		<tr>
			<td>Dipotassium Phosphate</td>
			<td>VWR</td>
			<td>BDH9266</td>
			<td>&#8805;98 %</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sigma Aldrich</td>
			<td>G5516</td>
			<td>&#8805;99.5 %</td>
		</tr>
		<tr>
			<td>Heraus Megafuge 16R Centrifuge</td>
			<td>Thermo Fischer Scientific</td>
			<td>750004271</td>
			<td>Maximum RCF 25,830 x g</td>
		</tr>
		<tr>
			<td>HR-MAS Disposable Insert Kit</td>
			<td>Bruker</td>
			<td>B4493</td>
			<td>Kel-F</td>
		</tr>
		<tr>
			<td>Iron(II) Sulfate Heptahydrate</td>
			<td>Alfa Aesar</td>
			<td>14498</td>
			<td>&#8805;99+ %</td>
		</tr>
		<tr>
			<td>Magnesium Sulfate Heptahydrate</td>
			<td>VWR</td>
			<td>10034998</td>
			<td>store at 18-26 &#176;C</td>
		</tr>
		<tr>
			<td>Manganese(II) Chloride Tetrahydrate</td>
			<td>Alfa Aesar</td>
			<td>11563</td>
			<td>&#8805;99 %</td>
		</tr>
		<tr>
			<td>Monopotassium Phosphate</td>
			<td>VWR</td>
			<td>470302-254</td>
			<td>&#8805;99 %</td>
		</tr>
		<tr>
			<td>pH Meter</td>
			<td>Mettler Toledo</td>
			<td>B706689216</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetrasodium Ethylenediaminetetraacetate</td>
			<td>Acros Organics</td>
			<td>13235-36-9</td>
			<td>&#8805;99.5 %</td>
		</tr>
		<tr>
			<td>Zinc Sulfate Heptahydrate</td>
			<td>Alfa Aesar</td>
			<td>33399</td>
			<td>&#8805;98 %</td>
		</tr>
		<tr>
			<td>12C3, d8-glycerol</td>
			<td>Cambridge Isotope Laboratory</td>
			<td>CDLM-8660</td>
			<td>12C3, 99.95%; D8, 98%</td>
		</tr>
		<tr>
			<td>13C6-glucose</td>
			<td>Sigma Alrdrich</td>
			<td>364606</td>
			<td>&#8805;99 % (CP)</td>
		</tr>
		<tr>
			<td>15N-sodium nitrate</td>
			<td>Sigma Aldrich</td>
			<td>364606</td>
			<td>&#8805;98 % 15N, &#8805;99 (cp)</td>
		</tr>
		<tr>
			<td>3.2 mm sapphire NMR rotor</td>
			<td>Cortecnet</td>
			<td>B6939</td>
			<td></td>
		</tr>
		<tr>
			<td>3.2 mm Silicone plug</td>
			<td>Bruker</td>
			<td>B7089</td>
			<td></td>
		</tr>
		<tr>
			<td>4 mm MAS Rotor Kit</td>
			<td>Bruker</td>
			<td>H14355</td>
			<td>Zirconia</td>
		</tr>
	</tbody>
</table>

preparation of fungal and plant materials for structural elucidation using dynamic nuclear polarization solid-state nmr

This protocol shows how uniformly 13C, 15N-labeled fungal materials can be produced and how these soft materials should be proceeded for solid-state NMR and sensitivity-enhanced DNP experiments. The sample processing procedure of plant biomass is also detailed. This method allows the measurement of a series of 1D and 2D 13C-13C/15N correlations spectra, which enables high-resolution structural elucidation of complex biomaterials in their native state, with minimal perturbation. The isotope-labeling can be examined by quantifying the intensity in 1D spectra and the polarization transfer efficiency in 2D correlation spectra. The success of dynamic nuclear polarization (DNP) sample preparation can be evaluated by the sensitivity enhancement factor. Further experiments examining the structural aspects of the polysaccharides and proteins will lead to a model of the three-dimensional architecture. These methods can be modified and adapted to investigate a wide range of carbohydrate-rich materials, including the natural cell walls of plants, fungi, algae and bacteria, as well as synthesized or designed carbohydrate polymers and their complex with other molecules.

The isotope labeling substantially enhances the NMR sensitivity and makes it possible for measuring a series of 2D 13C-13C and 13C-15N correlation spectra to analyze the composition, hydration, mobility and packing of polymers, which will be integrated to construct a three-dimensional model of cell wall architecture (Figure 1). If the uniform labeling succeeds, a complete set of 1D 13C and 15N ...

Watch this Scientific Journal Video about Preparation of Fungal and Plant Materials for Structural Elucidation Using Dynamic Nuclear Polarization Solid-State NMR at JoVE.com

Preparation of Fungal and Plant Materials for Structural Elucidation Using Dynamic Nuclear Polarization Solid-State NMR

Preparação de fungos e materiais vegetais para a elucidação estrutural usando polarização Nuclear dinâmica NMR Solid-State

A protocol for preparing 13C,15N-labeled fungal and plant samples for multidimensional solid-state NMR spectroscopy and dynamic nuclear polarization (DNP) investigations is presented.

This protocol shows how uniformly 13C, 15N-labeled fungal materials can be produced and how these soft materials should be proceeded for solid-state NMR ...

preparation-fungal-plant-materials-for-structural-elucidation-using

Research

JoVE Journal

Biochemistry

7.4K visualizações.  Louisiana State University. Este protocolo pode ser usado para preparar materiais fúngicos e vegetais para ressonância magnética nuclear de estado sólido, ou RN, e polarização nuclear dinâmica, ou DNP, experimentos para caracterização complexa do biossistema. Essa técnica nos permite investigar biomateriais a um nível de resolução atômica em seu ambiente nativo, por exemplo, em células inteiras. A aquisição de informações estruturais de alta resolução sobre materiais fúngicos ajudará no desenvolvi...

Vídeo: Preparation of Fungal and Plant Materials for Structural Elucidation Using Dynamic Nuclear Polarization Solid-State NMR

Preparation and Delivery of Protein Microcrystals in Lipidic Cubic Phase for Serial Femtosecond Crystallography

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural information, typically obtained by crystallography; however MPs are difficult to crystallize. Recent progress in MP structural determination has benefited greatly from the development of lipidic cubic phase (LCP) crystallization methods, which typically yield well-diffracting, but often small crystals that suffer from radiation damage during traditional crystallographic data collection at synchrotron sources. The development of new-generation X-ray free-electron laser (XFEL) sources that produce extremely bright femtosecond pulses has enabled room temperature data collection from microcrystals with no or negligible radiation damage. Our recent efforts in combining LCP technology with serial femtosecond crystallography (LCP-SFX) have resulted in high-resolution structures of several human G protein-coupled receptors, which represent a notoriously difficult target for structure determination. In the LCP-SFX technique, LCP is recruited as a matrix for both growth and delivery of MP microcrystals to the intersection of the injector stream with an XFEL beam for crystallographic data collection. It has been demonstrated that LCP-SFX can substantially improve the diffraction resolution when only sub-10 &#181;m crystals are available, or when the use of smaller crystals at room temperature can overcome various problems associated with larger cryocooled crystals, such as accumulation of defects, high mosaicity and cryocooling artifacts. Future advancements in X-ray sources and detector technologies should make serial crystallography highly attractive and practicable for implementation not only at XFELs, but also at more accessible synchrotron beamlines. Here we present detailed visual protocols for the preparation, characterization and delivery of microcrystals in LCP for serial crystallography experiments. These protocols include methods for conducting crystallization experiments in syringes, detecting and characterizing the crystal samples, optimizing crystal density, loading microcrystal laden LCP into the injector device and delivering the sample to the beam for data collection.

We describe procedures for the preparation and delivery of membrane protein microcrystals in lipidic cubic phase for serial crystallography at X-ray free-electron lasers and synchrotron sources. These protocols can also be applied for incorporation and delivery of soluble protein microcrystals, leading to substantially reduced sample consumption compared to liquid injection.

Preparação e entrega de microcristais de proteína na fase cúbica lipídico para Serial Femtosecond Cristalografia

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural ...

Bioquímica

A Protein Preparation Method for the High-throughput Identification of Proteins Interacting with a Nuclear Cofactor Using LC-MS/MS Analysis

Transcriptional coregulators are vital to the efficient transcriptional regulation of nuclear chromatin structure. Coregulators play a variety of roles in regulating transcription. These include the direct interaction with transcription factors, the covalent modification of histones and other proteins, and the occasional chromatin conformation alteration. Accordingly, establishing relatively quick methods for identifying proteins that interact within this network is crucial to enhancing our understanding of the underlying regulatory mechanisms. LC-MS/MS-mediated protein binding partner identification is a validated technique used to analyze protein-protein interactions. By immunoprecipitating a previously-identified member of a protein complex with an antibody (occasionally with an antibody for a tagged protein), it is possible to identify its unknown protein interactions via mass spectrometry analysis. Here, we present a method of protein preparation for the LC-MS/MS-mediated high-throughput identification of protein interactions involving nuclear cofactors and their binding partners. This method allows for a better understanding of the transcriptional regulatory mechanisms of the targeted nuclear factors.

We have established a method for the purification of coregulatory interaction proteins using the LC-MS/MS system.

Uma proteína Método de Preparação para a Identificação de alto rendimento de proteínas que interagem com um co-fator nuclear utilizando LC-MS / MS Análise

Transcriptional coregulators are vital to the efficient transcriptional regulation of nuclear chromatin structure. Coregulators play a variety of roles in ...

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the propagation of neurodegenerative disorders. Such assemblies consist in multiple protein subunits organized in a non-covalent way to form large macromolecular objects that can execute a variety of cellular functions or cause detrimental consequences. Atomic insights into the assembly mechanisms and the functioning of those macromolecular assemblies remain often scarce since their inherent insolubility and non-crystallinity often drastically reduces the quality of the data obtained from most techniques used in structural biology, such as X-ray crystallography and solution Nuclear Magnetic Resonance (NMR). We here present magic-angle spinning solid-state NMR spectroscopy (SSNMR) as a powerful method to investigate structures of macromolecular assemblies at atomic resolution. SSNMR can reveal atomic details on the assembled complex without size and solubility limitations. The protocol presented here describes the essential steps from the production of 13C/15N isotope-labeled macromolecular protein assemblies to the acquisition of standard SSNMR spectra and their analysis and interpretation. As an example, we show the pipeline of a SSNMR structural analysis of a filamentous protein assembly.

Structures of supramolecular protein assemblies at atomic resolution are of high relevance because of their crucial roles in a variety of biological phenomena. Herein, we present a protocol to perform high-resolution structural studies on insoluble and non-crystalline macromolecular protein assemblies by magic-angle spinning solid-state nuclear magnetic resonance spectroscopy (MAS SSNMR).

Escala atômica estudos estruturais de conjuntos macromoleculares por espectroscopia de estado sólido da ressonância magnética Nuclear

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the ...

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to precisely maneuver the nucleation and crystal growth towards desired sizes of protein crystals. The controlled crystallization is based on sample investigation with in situ Dynamic Light Scattering (DLS) while all visual changes in the droplet are monitored online with the help of a microscope coupled to a CCD camera, thus enabling a full investigation of the protein droplet during all stages of crystallization. The use of in situ DLS measurements throughout the entire experiment allows a precise identification of the highly supersaturated protein solution transitioning to a new phase &#8211; the formation of crystal nuclei. By identifying the protein nucleation stage, the crystallization can be optimized from large protein crystals to the production of protein microcrystals. The experimental protocol shows an interactive crystallization approach based on precise automated steps such as precipitant addition, water evaporation for inducing high supersaturation, and sample dilution for slowing induced homogeneous nucleation or reversing phase transitions.

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

Here we present a protocol for controlled production of protein microcrystals. The process uses an automated device allowing controlled manipulation of several crystallization parameters. The protein crystallization is carried-out by controlled and automated addition of crystallization solutions while monitoring and investigating the radius distribution of particles in the crystallization droplet.

Crescimento de cristais de proteína com dimensões distintas usando automatizado cristalização juntamente com In Situ Difusão dinâmica da luz

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to ...

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be employed using a conventional confocal microscope equipped with digital detectors. A protocol for the use of the technique in vitro is shown by means of a use case where number and brightness can be seen to accurately quantify the oligomeric state of mVenus-labelled FKBP12F36V before and after the addition of the dimerizing drug AP20187. The importance of using the correct microscope acquisition parameters and the correct data preprocessing and analysis methods are discussed. In particular, the importance of the choice of photobleaching correction is stressed. This inexpensive method can be employed to study protein-protein interactions in many biological contexts.

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

This protocol describes a calibration-free approach for quantifying protein homo-oligomerization in vitro based on fluorescence fluctuation spectroscopy using commercial light scanning microscopy. The correct acquisition settings and analysis methods are shown.

Livre de calibração em Vitro quantificação da proteína Homo-oligomerização usando instrumentação comercial e livre, Open Source Software de análise de brilho

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be ...

Modifying Baculovirus Expression Vectors to Produce Secreted Plant Proteins in Insect Cells

It has been a challenge for scientists to express recombinant secretory eukaryotic proteins for structural and biochemical studies. The baculovirus-mediated insect cell expression system is one of the systems used to express recombinant eukaryotic secretory proteins with some post-translational modifications. The secretory proteins need to be routed through the secretory pathways for protein glycosylation, disulfide bonds formation, and other post-translational modifications. To improve the existing insect cell expression of secretory plant proteins, a baculovirus expression vector is modified by the addition of either a GP67 or a hemolin signal peptide sequence between the promoter and multiple-cloning sites. This newly designed modified vector system successfully produced a high yield of soluble recombinant secreted plant receptor proteins of Arabidopsis thaliana. Two of the expressed plant proteins, the extracellular domains of Arabidopsis TDR and PRK3 plasma membrane receptors, were crystallized for X-ray crystallographic studies. The modified vector system is an improved tool that can potentially be used for the expression of recombinant secretory proteins in the animal kingdom as well.

Here, we present the protocols for utilizing the insect cell and baculovirus protein expression system to produce large quantities of plant secreted proteins for protein crystallization. A baculovirus expression vector has been modified with either GP67 or insect hemolin signal peptide for plant protein secretion expression in insect cells.

Modificar vetores de expressão de Baculovirus para produzir secretado proteínas vegetais em células de inseto

It has been a challenge for scientists to express recombinant secretory eukaryotic proteins for structural and biochemical studies. The ...

Measuring Interactions of Globular and Filamentous Proteins by Nuclear Magnetic Resonance Spectroscopy (NMR) and Microscale Thermophoresis (MST)

Filamentous proteins such as vimentin provide organization within cells by providing a structural scaffold with sites that bind proteins containing plakin repeats. Here, a protocol for detecting and measuring such interactions is described using the globular plakin repeat domain of envoplakin and the helical coil of vimentin. This provides a basis for determining whether a protein binds vimentin (or similar filamentous proteins) and for measurement of the affinity of the interaction. The globular protein of interest is labeled with 15N and titrated with vimentin protein in solution. A two-dimensional NMR spectrum is acquired to detect interactions by observing changes in peak shape or chemical shifts, and to elucidate effects of solution conditions including salt levels, which influence vimentin quaternary structure. If the protein of interest binds the filamentous ligand, the binding interaction is quantified by MST using the purified proteins. The approach is a straightforward way for determining whether a protein of interest binds a filament, and for assessing how alterations, such as mutations or solution conditions, affect the interaction.

Measuring Interactions of Globular and Filamentous Proteins by Nuclear Magnetic Resonance Spectroscopy NMR and Microscale Thermophoresis MST

Here, we present a protocol for the production and purification of proteins that are labeled with stable isotopes, and subsequent characterization of protein-protein interactions using Nuclear Magnetic Resonance (NMR) spectroscopy and MicroScale Thermophoresis (MST) experiments.

Interações de proteínas globulares e filamentosas por espectroscopia de ressonância magnética Nuclear (NMR) e Thermophoresis de microescala (MST) de medição

Filamentous proteins such as vimentin provide organization within cells by providing a structural scaffold with sites that bind proteins containing plakin ...

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures and information on more challenging systems. Sample preparation for cryo-EM is undergoing a similar revolution with new approaches being developed to supersede the traditional blotting systems. These include the use of piezo-electric dispensers, pin printing and direct spraying. As a result of these developments, the speed of grid preparation is going from seconds to milliseconds, providing new opportunities, especially in the field of time-resolved cryo-EM where proteins and substrates can be rapidly mixed before plunge freezing, trapping short lived intermediate states. Here we describe, in detail, a standard protocol for making grids on our in-house time-resolved EM device both for standard fast grid preparation and also for time-resolved experiments. The protocol requires a minimum of about 50 &#181;L sample at concentrations of &#8805; 2 mg/mL for the preparation of 4 grids. The delay between sample application and freezing can be as low as 10 ms. One limitation is increased ice thickness at faster speeds and compared to the blotting method. We hope this protocol will aid others in designing their own grid making devices and those interested in designing time-resolved experiments.

Here, we provide a detailed protocol for the use of a rapid grid making device for both fast grid-making and for rapid mixing and freezing to conduct time-resolved experiments.

Preparação rápida da grade para microscopia crio-elétron resolvida pelo tempo

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures ...

Preparation of Sample Support Films in Transmission Electron Microscopy using a Support Floatation Block

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant bottleneck. Macromolecular samples are ideally imaged directly from random orientations in a thin layer of vitreous ice. However, many samples are refractory to this, and protein denaturation at the air-water interface is a common problem. To overcome such issues, support films-including amorphous carbon, graphene, and graphene oxide-can be applied to the grid to provide a surface which samples can populate, reducing the probability of particles experiencing the deleterious effects of the air-water interface. The application of these delicate supports to grids, however, requires careful handling to prevent breakage, airborne contamination, or extensive washing and cleaning steps. A recent report describes the development of an easy-to-use floatation block that facilitates wetted transfer of support films directly to the sample. Use of the block minimizes the number of manual handling steps required, preserving the physical integrity of the support film, and the time over which hydrophobic contamination can accrue, ensuring that a thin film of ice can still be generated. This paper provides step-by-step protocols for the preparation of carbon, graphene, and graphene oxide supports for EM studies.

Sample preparation for cryo-electron microscopy (cryo-EM) is a significant bottleneck in the structure determination workflow of this method. Here, we provide detailed methods for using an easy-to-use, three-dimensionally printed block for the preparation of support films to stabilize samples for transmission EM studies.

Preparação de filmes de suporte à amostra em microscopia eletrônica de transmissão usando um bloco de flutuação de suporte

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant ...

Pure Shift Nuclear Magnetic Resonance: a New Tool for Plant Metabolomics

Nuclear Magnetic Resonance (NMR) is one of the most powerful tools used in metabolomics. It stands as a highly accurate and reproducible method that not only provides quantitative data but also permits structural identification of the metabolites present in complex mixtures.
Metabolic profiling by 1H NMR has proven useful in the study of various types of plant scenarios, which include the evaluation of crop&#160;conditions, harvest and post- harvest treatments, metabolic phenotyping, metabolic pathways, gene regulation, identification of biomarkers, chemotaxonomy, quality control, denomination of origin, among others. However, signal overlapping of the large number of resonances with expanded J-coupling multiplicities complicates the spectra analysis and its interpretation, and represents a limitation for classical 1H NMR profiling.
In the last decade, novel NMR broadband homonuclear decoupling techniques through which multiplet signals collapse into single resonance lines - commonly called Pure Shift methods - have been developed to overcome the spectra resolution problem inherent to 1H NMR classical spectra.
Here a step-by-step protocol of the plant extract preparation and the procedure to record optimal Pure Shift PSYCHE and SAPPHIRE-PSYCHE spectra in three different plant matrices - Vanilla plant&#160;leaves, potato tubers (S. tuberosum), and Cape gooseberries (P. peruviana) - is presented. The effect of the gain in resolution in metabolic identification, correlation analysis and multivariate analyses, as compared against classical spectra, is discussed.

<meta charset="utf8"></head><body>Ressonância magnética nuclear de deslocamento puro: uma nova ferramenta para metabolômica de plantas

This paper presents the use of PSYCHE and SAPPHIRE-PSYCHE in the metabolic profiling of plants and includes detailed procedures for sample preparation and optimal Pure Shift NMR spectra recording. Examples through which the gain in resolution achieved by homonuclear decoupling allows a more comprehensive understanding of the system are discussed.

Ressonância magnética nuclear de deslocamento puro: uma nova ferramenta para metabolômica de plantas

Nuclear Magnetic Resonance (NMR) is one of the most powerful tools used in metabolomics. It stands as a highly accurate and reproducible method that not ...