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 &amp;deg;C</td></tr><tr><td>Cobalt(II) Chloride Hexahydrate</td><td>Honeywell|Fluka</td><td>60820</td><td>&amp;ge;98 %</td></tr><tr><td>Copper(II) Sulfate Pentahydrate</td><td>BDH</td><td>BDH9312</td><td>&amp;ge;98 %</td></tr><tr><td>Corning LSE shaking incubator</td><td>Thermo Fisher Scientific</td><td>7202152</td><td></td></tr><tr><td>D&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma Aldrich</td><td>151882</td><td>99.9 atom % D</td></tr><tr><td>d&lt;sub&gt;6&lt;/sub&gt;-DMSO</td><td>Sigma Aldrich</td><td>151874</td><td>99.9 atom % D</td></tr><tr><td>d&lt;sub&gt;8&lt;/sub&gt;-glycerol</td><td>Sigma Aldrich</td><td>447498</td><td>&amp;ge;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>&amp;ge;98 %</td></tr><tr><td>Glycerol</td><td>Sigma Aldrich</td><td>G5516</td><td>&amp;ge;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>&amp;ge;99+ %</td></tr><tr><td>Magnesium Sulfate Heptahydrate</td><td>VWR</td><td>10034998</td><td>store at 18-26 &amp;deg;C</td></tr><tr><td>Manganese(II) Chloride Tetrahydrate</td><td>Alfa Aesar</td><td>11563</td><td>&amp;ge;99 %</td></tr><tr><td>Monopotassium Phosphate</td><td>VWR</td><td>470302-254</td><td>&amp;ge;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>&amp;ge;99.5 %</td></tr><tr><td>Zinc Sulfate Heptahydrate</td><td>Alfa Aesar</td><td>33399</td><td>&amp;ge;98 %</td></tr><tr><td>&lt;sup&gt;12&lt;/sup&gt;C3, d&lt;sub&gt;8&lt;/sub&gt;-glycerol</td><td>Cambridge Isotope Laboratory</td><td>CDLM-8660</td><td>&lt;sup&gt;12&lt;/sup&gt;C3, 99.95%; D8, 98%</td></tr><tr><td>&lt;sup&gt;13&lt;/sup&gt;C6-glucose</td><td>Sigma Alrdrich</td><td>364606</td><td>&amp;ge;99 % (CP)</td></tr><tr><td>&lt;sup&gt;15&lt;/sup&gt;N-sodium nitrate</td><td>Sigma Aldrich</td><td>364606</td><td>&amp;ge;98 % 15N, &amp;ge;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

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 Views.  Louisiana State University. Dit protocol kan worden gebruikt om schimmel- en plantenmaterialen voor te bereiden op vaste nucleaire magnetische resonantie, of NMR, en dynamische nucleaire polarisatie, of DNP, experimenten voor complexe biosysteemkarakterisering. Deze techniek stelt ons in staat om biomaterialen te onderzoeken tot een atomair resolutieniveau in hun eigen omgeving, bijvoorbeeld in hele cellen. De verwerving van structurele informatie over schimmelmaterialen met hoge resolutie zal helpen bij de ontwikkeling...

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

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).

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

Dissolution Dynamic Nuclear Polarization Instrumentation for Real-time Enzymatic Reaction Rate Measurements by NMR

The main limitation of NMR-based investigations is low sensitivity. This prompts for long acquisition times, thus preventing real-time NMR measurements of metabolic transformations. Hyperpolarization via dissolution DNP circumvents part of the sensitivity issues thanks to the large out-of-equilibrium nuclear magnetization stemming from the electron-to-nucleus spin polarization transfer. The high NMR signal obtained can be used to monitor chemical reactions in real time. The downside of hyperpolarized NMR resides in the limited time window available for signal acquisition, which is usually on the order of the nuclear spin longitudinal relaxation time constant, T1, or, in favorable cases, on the order of the relaxation time constant associated with the singlet-state of coupled nuclei, TLLS. Cellular uptake of endogenous molecules and metabolic rates can provide essential information on tumor development and drug response. Numerous previous hyperpolarized NMR studies have demonstrated the relevancy of pyruvate as a metabolic substrate for monitoring enzymatic activity in vivo. This work provides a detailed description of the experimental setup and methods required for the study of enzymatic reactions, in particular the pyruvate-to-lactate conversion rate in presence of lactate dehydrogenase (LDH), by hyperpolarized NMR.

The sensitivity enhancement provided by dissolution dynamic nuclear polarization (DNP) enables following metabolic processes in real time by NMR and MRI. The characteristics and performances of a dedicated dissolution DNP setup designed for study enzymatic reactions are discussed.

The main limitation of NMR-based investigations is low sensitivity. This prompts for long acquisition times, thus preventing real-time NMR measurements of ...

Chemistry

Sequencing of Plant Wall Heteroxylans Using Enzymic, Chemical (Methylation) and Physical (Mass Spectrometry, Nuclear Magnetic Resonance) Techniques

This protocol describes the specific techniques used for the characterization of reducing end (RE) and internal region glycosyl sequence(s) of heteroxylans. De-starched wheat endosperm cell walls were isolated as an alcohol-insoluble residue (AIR)1 and sequentially extracted with water (W-sol Fr) and 1 M KOH containing 1% NaBH4 (KOH-sol Fr) as described by Ratnayake et al. (2014)2. Two different approaches (see summary in Figure 1) are adopted. In the first, intact W-sol AXs are treated with 2AB to tag the original RE backbone chain sugar residue and then treated with an endoxylanase to generate a mixture of 2AB-labelled RE and internal region reducing oligosaccharides, respectively. In a second approach, the KOH-sol Fr is hydrolyzed with endoxylanase to first generate a mixture of oligosaccharides which are subsequently labelled with 2AB. The enzymically released ((un)tagged) oligosaccharides from both W- and KOH-sol Frs are then methylated and the detailed structural analysis of both the native and methylated oligosaccharides is performed using a combination of MALDI-TOF-MS, RP-HPLC-ESI-QTOF-MS and ESI-MSn. Endoxylanase digested KOH-sol AXs are also characterized by nuclear magnetic resonance (NMR) that also provides information on the anomeric configuration. These techniques can be applied to other classes of polysaccharides using the appropriate endo-hydrolases.

Sequencing of Plant Wall Heteroxylans Using Enzymic, Chemical Methylation and Physical Mass Spectrometry, Nuclear Magnetic Resonance Techniques

This protocol describes the specific techniques used for the structural characterization of reducing end (RE) and internal region glycosyl sequence(s) of heteroxylans by tagging the RE with 2 aminobenzamide prior to enzymatic (endoxylanase) hydrolysis and then analysis of the resultant oligosaccharides using mass spectrometry (MS) and nuclear magnetic resonance (NMR).

This protocol describes the specific techniques used for the characterization of reducing end (RE) and internal region glycosyl sequence(s) of ...

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.

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.

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 ...

Paramagnetic Relaxation Enhancement for Detecting and Characterizing Self-Associations of Intrinsically Disordered Proteins

Intrinsically disordered proteins and intrinsically disordered regions within proteins make up a large and functionally significant part of the human proteome. The highly flexible nature of these sequences allows them to form weak, long-range, and transient interactions with diverse biomolecular partners. Specific yet low-affinity interactions promote promiscuous binding and enable a single intrinsically disordered segment to interact with a multitude of target sites. Because of the transient nature of these interactions, they can be difficult to characterize by structural biology methods that rely on proteins to form a single, predominant conformation. Paramagnetic relaxation enhancement NMR is a useful tool for identifying and defining the structural underpinning of weak and transient interactions. A detailed protocol for using paramagnetic relaxation enhancement to characterize the lowly-populated encounter complexes that form between intrinsically disordered proteins and their protein, nucleic acid, or other biomolecular partners is described.

A protocol for the application of paramagnetic relaxation enhancement NMR spectroscopy to detect weak and transient inter- and intra-molecular interactions in intrinsically disordered proteins is presented.

Intrinsically disordered proteins and intrinsically disordered regions within proteins make up a large and functionally significant part of the human ...

High-Resolution Studies of Proteins in Natural Membranes by Solid-State NMR

Membrane proteins are vital for cell function and thus represent important drug targets. Solid-state Nuclear Magnetic Resonance (ssNMR) spectroscopy offers a unique access to probe the structure and dynamics of such proteins in biological membranes of increasing complexity. Here, we present modern solid-state NMR spectroscopy as a tool to study structure and dynamics of proteins in natural lipid membranes and at atomic scale. Such spectroscopic studies profit from the use of high-sensitivity ssNMR methods, i.e., proton-(1H)-detected ssNMR and DNP (Dynamic Nuclear Polarization) supported ssNMR. Using bacterial outer membrane beta-barrel protein BamA and the ion channel KcsA, we present methods to prepare isotope-labeled membrane proteins and to derive structural and motional information by ssNMR.

This work details robust basic routines on how to prepare isotope-labeled membrane protein samples and analyze them at high-resolution with modern solid-state NMR spectroscopy methods.

Membrane proteins are vital for cell function and thus represent important drug targets. Solid-state Nuclear Magnetic Resonance (ssNMR) spectroscopy ...

Quantitative 31P NMR Analysis of Lignins and Tannins

The development of sustainable biorefinery products is confronted, among others, with the challenge of lignin and tannin valorization. These abundant, renewable aromatic biopolymers have not been widely exploited due to their inherent structural complexity and high degrees of variability and species diversity. The lack of a defined primary structure for these polyphenols is further compounded with complex chemical alterations induced during processing, eventually imparting a large variety of structural features of extreme significance for any further utilization efforts.
Consequently, a protocol for the rapid, simple, and unequivocal identification and quantification of the various functional groups present in natural polyphenols, is a fundamental prerequisite for understanding and accordingly tailor their reactivity and eventual utility.
Quantitative 31P NMR offers the opportunity to rapidly and reliably identify unsubstituted, o-mono substituted, and o-disubstituted phenols, aliphatic OHs, and carboxylic acid moieties in lignins and tannins with broad application potential.
The methodology consists of an in situ quantitative lignin or tannin labeling procedure using a suitable 31P containing probe, followed by the acquisition of a quantitative 31P NMR spectrum in the presence of an internal standard. The high natural abundance of the 31P nucleus allows for small amounts of the sample (~30 mg) and short NMR acquisition times (~30-120 min) with well-resolved 31P signals that are highly dependent on the surrounding chemical environment of the labeled OH groups.

Quantitative 31P NMR Analysis of Lignins and Tannins

31P NMR is a powerful tool for the structural elucidation of polyphenols. This fast, easy, precise, quantitative, and highly reproducible analytical procedure, that allows for the quantification and differentiation of the different types of hydroxy, phenolic, and carboxylic groups in lignins and tannins has now become a routine analytical tool.

The development of sustainable biorefinery products is confronted, among others, with the challenge of lignin and tannin valorization. These abundant, ...

Protocol for NMR 15N Relaxation and hetNOE Experiments in Protein Dynamics Study

NMR 15N Relaxation Experiments for the Investigation of Picosecond to Nanoseconds Structural Dynamics of Proteins

Nuclear magnetic resonance (NMR) spectroscopy allows studying proteins in solution and under physiological temperatures. Frequently, either the amide groups of the protein backbone or the methyl groups in side chains are used as reporters of structural dynamics in proteins. A structural dynamics study of the protein backbone of globular proteins on 15N labeled and fully protonated samples usually works well for proteins with a molecular weight of up to 50 kDa. When side chain deuteration in combination with transverse relaxation optimized spectroscopy (TROSY) is applied, this limit can be extended up to 200 kDa for globular proteins and up to 1 MDa when the focus is on the side chains. When intrinsically disordered proteins (IDPs) or proteins with intrinsically disordered regions (IDRs) are investigated, these weight limitations do not apply but can go well beyond. The reason is that IDPs or IDRs, characterized by high internal flexibility, are frequently dynamically decoupled. Various NMR methods offer atomic-resolution insights into structural protein dynamics across a wide range of time scales, from picoseconds up to hours. Standard 15N relaxation measurements overview a protein&#39;s internal flexibility and characterize the protein backbone dynamics experienced on the fast pico- to nanosecond timescale. This article presents a hands-on protocol for setting up and recording NMR 15N R1, R2, and heteronuclear Overhauser effect (hetNOE) experiments. We show exemplary data and explain how to interpret them simply qualitatively before any more sophisticated analysis.

Author Spotlight: Exploring Intrinsically Disordered Protein Dynamics Through NMR Relaxation Experiments

Nuclear magnetic resonance (NMR) spectroscopy can characterize structural protein dynamics in a residue-specific manner. We provide a hands-on protocol for recording NMR 15N R1 and R2 relaxation and {1H}-15N heteronuclear Overhauser effect (hetNOE) experiments, sensitive to the picoseconds to nanoseconds timescale.

Nuclear magnetic resonance (NMR) spectroscopy allows studying proteins in solution and under physiological temperatures. Frequently, either the amide ...

Isolation and Biophysical Study of Fruit Cuticles

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and abiotic stresses and also regulates water flow from the external environment.1 A biopolyester (cutin) and long-chain fatty acids (waxes) form the principal structural framework of the cuticle; the functional integrity of the cuticular layer depends on the outer 'epicuticular' layer as well as the blend consisting of the cutin biopolymer and 'intracuticular' waxes.2 Herein, we describe a comprehensive protocol to extract waxes exhaustively from commercial tomato (Solanum lycopersicum) fruit cuticles or to remove epicuticular and intracuticular waxes sequentially and selectively from the cuticle composite. The method of Jetter and Sch&auml;ffer (2001) was adapted for the stepwise extraction of epicuticular and intracuticular waxes from the fruit cuticle.3,4 To monitor the process of sequential wax removal, solid-state cross-polarization magic-angle-spinning (CPMAS) 13C NMR spectroscopy was used in parallel with atomic force microscopy (AFM), providing molecular-level structural profiles of the bulk materials complemented by information on the microscale topography and roughness of the cuticular surfaces. To evaluate the cross-linking capabilities of dewaxed cuticles from cultivated wild-type and single-gene mutant tomato fruits, MAS 13C NMR was used to compare the relative proportions of oxygenated aliphatic (CHO and CH2O) chemical moieties.
Exhaustive dewaxing by stepwise Soxhlet extraction with a panel of solvents of varying polarity provides an effective means to isolate wax moieties based on the hydrophobic characteristics of their aliphatic and aromatic constituents, while preserving the chemical structure of the cutin biopolyester. The mechanical extraction of epicuticular waxes and selective removal of intracuticular waxes, when monitored by complementary physical methodologies, provides an unprecedented means to investigate the cuticle assembly: this approach reveals the supramolecular organization and structural integration of various types of waxes, the architecture of the cutin-wax matrix, and the chemical composition of each constituent. In addition, solid-state 13C NMR reveals differences in the relative numbers of CHO and CH2O chemical moieties for wild-type and mutant red ripe tomato fruits. The NMR techniques offer exceptional tools to fingerprint the molecular structure of cuticular materials that are insoluble, amorphous, and chemically heterogeneous. As a noninvasive surface-selective imaging technique, AFM furnishes an effective and direct means to probe the structural organization of the cuticular assembly on the nm-&mu;m length scale.

Aerial plant organs are protected by the cuticle, a supramolecular biopolyester-wax assembly. We present protocols to monitor selective removal of epi- and intracuticular waxes from tomato fruit cuticles on molecular and micro scales by solid-state NMR and atomic force microscopy, respectively, and to assess the cross-linking capacity of engineered cuticular biopolyesters.

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and ...

Biology

Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability

Dynamic nuclear polarization (DNP) can dramatically increase the sensitivity of magic angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy. These sensitivity gains increase as temperatures decrease and are large enough to enable the study of molecules at very low concentrations at the operating temperatures (~100 K) of most commercial DNP-equipped NMR spectrometers. This leads to the possibility of in-cell structural biology on cryopreserved cells for macromolecules at their endogenous levels in their native environments. However, the freezing rates required for cellular cryopreservation are exceeded during typical sample handling for DNP MAS NMR and this results in loss of cellular integrity and viability. This article describes a detailed protocol for the preparation and cryogenic transfer of a frozen sample of mammalian cells into a MAS NMR spectrometer.

Presented here is a protocol for cryogenic transfer of frozen samples into the dynamic nuclear polarization (DNP) magic angle spinning (MAS) nuclear magnetic resonance (NMR) probe. The protocol includes directions for rotor storage prior to the experiment and directions for viability measurements before and after the experiment.

Dynamic nuclear polarization (DNP) can dramatically increase the sensitivity of magic angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy. ...

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

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