The authors would like to acknowledge Genome Canada and Genome British Columbia for platform funding, and support. We also thank Dr. Carol E. Parker for critical review of the manuscript and editing assistance. CHL also thanks the British Columbia Proteomics Network for support.

1. Tissue Sectioning
<ol>
	<li>Flash-freeze the issue specimens, once harvested, using liquid nitrogen, ship them on dry ice (if shipping is required), and store them at -80 &#176;C until tissue sectioning. (If commercial samples are used, ensure that the samples are prepared in this manner.)</li>
	<li>Cut organs to a manageable size to fit the MALDI target. Trim off any unwanted parts of the organ. For this study described here, bovine calf lenses were decapsulated using a previously-described procedure35 before tissue sectioning.</li>
	<li>Remove whole organs from the -80 &#176;C freezer and fix them onto a cryogenic tissue cutting stage. To fix a bovine calf lens, place one or two drops of water on the tissue-cutting stage of a&#160;cryostat. Quickly place the lens in the water before it solidifies. Alternatively, optimal cutting temperature (OCT) compounds can also be used to fix a tissue onto the cutting stage. If OCT compounds are used, a minimal amount of OCT should be applied and care should be taken to ensure that the cut tissue sections will not be contaminated with OCT compounds which can interfere with ionization and detection of the analytes5,36,37.</li>
	<li>Allow the temperature inside the cryostat to equilibrate to -18 &#176;C. Colder or warmer temperatures can be used for softer or harder tissues, respectively. Then cut the tissue equatorially into 20&#160;&#956;m thick slices. Use 10-15 &#956;m thick slices for most tissues; however, because of the fragile nature of the bovine lens tissue, 20&#160;&#956;m thick slices were used. For bovine lens tissue, discard the first tissue sections and only use slices which are close to or at the equatorial plane.</li>
	<li>If an ocular lens tissue is imaged, use 1.5 &#956;l of formic acid (98% in purity, LC-MS grade) to prewet the surface of an indium tin oxide (ITO)-coated glass slide.</li>
	<li>Carefully transfer the tissue sections to the surface of the ITO-coated glass microscopic slide inside the cryostat. The tissue section will quickly thaw and will become tightly affixed to the slide surface. Usually, multiple tissue sections can be mounted onto a same ITO-coated slide in this way.</li>
	<li>Lyophilize the slide&#160;for 15 min before MALDI matrix application.</li>
	<li>For matrix testing, dissolve the individual matrices in appropriate solvents. Manually spot 1 &#956;l of each matrix solution onto the tissue section. Additionally, spot a small-molecule calibration standard onto the tissue for verification of MALDI sensitivity.</li>
	<li>Add three teaching marks to the ITO coated glass slide&#160;by writing on the nonconductive surface of the ITO coated glass slide with a correction-fluid pen. Take an optical image of the tissue slide using a flatbed scanner and save it in an appropriate format such as tiff or jpg.</li>
</ol>
2. Matrix Coating
2.1. Automated Matrix Coating
<ol>
	<li>Apply matrix solutions, which contain acetonitrile or mixed acetonitrile/water as solvents, automatically to the surfaces of the tissue sections using a Bruker ImagePrep or a similar electronic matrix sprayer.</li>
	<li>Cover the edges of the front surface of the ITO-coated glass slide&#160;with tape so that the matrix does not coat the edges of the slide. This ensures that the teaching marks on the opposite surface can be used for tissue slide alignment. Do not cover the edges of the slide with matrix as they are used as contact points to maintain the electrical conductivity of the ITO-coated slide.</li>
	<li>Coat the glass slide by using twenty cycles of matrix coating (2-sec spray, 30-sec incubation, and 60-sec drying time for each cycle).</li>
</ol>
2.2. Manual Matrix Coating
<ol>
	<li>If organic solvents (e.g.&#160;chloroform and ethyl acetate) that are incompatible with the manufacturing materials of the electronic matrix sprayer are required, use a pneumatically-assisted airbrush sprayer to apply the matrix. Add the prepared matrix solution to the solvent reservoir of the airbrush gun and apply a gentle flow of pressurized nitrogen gas to prime the spray.</li>
	<li>Cover the edges of the front surface of the ITO-coated glass slides with tape so that the matrix does not coat the edges of the slide. This ensures that the teaching marks on the opposite surface can be used for tissue slide alignment. Do not cover the edges of the slide with matrix as they are used as contact points to maintain the electrical&#160;conductivity of the ITO-coated slide.</li>
	<li>After stable and fine sprays have been observed, manually spray the matrix so that it completely coats the tissue section. Apply the minimum amount of matrix solution required to barely wet the surface during each cycle to prevent possible analyte delocalization. In general, use approximately 10 cycles of matrix spray to coat a tissue section; the number of cycles is dependent on the tissue type and matrix composition.</li>
</ol>
3. MALDI MS
<ol>
	<li>Prepare a mass calibration solution by diluting the &#34;ES Tuning Mix&#34; standard solution by a factor of 1:200 in 60:40 isopropanol:water (containing 0.1% formic acid in the final mixture).</li>
	<li>Introduce 2 &#956;l/min of the diluted &#34;ES Tuning Mix&#34; solution into the dual-mode electrospray ionization (ESI)/MALDI ion source on the FTICR mass spectrometer, from the ESI side.</li>
	<li>Operate the FTICR instrument in the positive-ion ESI mode, with broadband detection and a data acquisition size of 1,024 kb/sec. Typical ESI parameters are capillary electrospray voltage, 3,900 V; spray shield voltage,&#160;3,600 V; nebulizer gas (N2) flow, 2 L/min;&#160;dry gas (N2) flow and temperature, 4 L/min and 200 &#176;C; skimmer 1 voltage, 15 V; time of flight (TOF), 0.009 sec; collision gas (Ar) flow, 0.4 L/s; source ion accumulation time, 0.1 sec; and collision cell ion accumulation time, 0.2 sec. Tune the FTICR operation parameters in order to maximize the analytical sensitivity over the mass range from m/z 200 to 1,400, while maintaining good time-domain free-induction decay (FID) signals. Typically, the ICR operation parameters are sidekick voltage, 8 V; sidekick offset voltage, 8 V; excitation amplification of 10; excitation pulse time, 0.01-0.015 sec; front trap plate voltage, 1.5 V; back trap plate voltage, 1.6 V and analyzer entrance voltage, -4 V. After a set of FTICR operation parameters has been determined, acquire the ESI mass spectra and calibrate the instrument using the reference masses of the standard compounds in the &#34;ES Tuning Mix&#34; solution.</li>
	<li>To tune the instrument for MALDI operation, dissolve several 1&#160;&#956;l aliquots of a mixed terfenadine and reserpine standard solution in the matrix solution at a concentration of 1 &#956;M each, and spot these solutions directly onto one of the tissue sections (i.e.&#160;a test tissue section) which has been mounted on an ITO-coated slide. Place the ITO-coated slide into a tissue slide adaptor (i.e.&#160;a special MALDI target) and load the adaptor from the MALDI side into the dual ESI/MALDI ion source. Optimize the appropriate MALDI operating parameters for the laser power and the number of laser shots for MALDI signal accumulation for each mass scan, etc. Typical MALDI operation parameters are: laser shots, 50; and a MALDI plate voltage of 300 V.</li>
	<li>After tuning, calibrating, and optimizing the instrument for MALDI-MSI experiments, align the physical location of a tissue section to be imaged with its recorded optical image within the imaging software. Use the three &#34;correction-fluid&#34; marks, which had been previously put on the opposite side of the ITO-coated slide surface (step 1.9), for this alignment using a three-point triangulation method.</li>
	<li>Perform a simultaneous ESI and MALDI operation so that each mass spectrum contains the reference mass peaks of the &#34;ES Tuning Mix&#34; solution for post-acquisition internal mass calibration. This will result in the most accurate mass measurement during MALDI MS. To do this, first attenuate the ESI signal by decreasing the capillary voltage until the MALDI signals dominate the spectra while the ESI calibrant signals are still high enough for internal mass calibration.</li>
	<li>Next, set up an automated rastering method for laser irradiation. Define the tissue regions to be imaged and set the appropriate laser raster step size. Note that smaller raster step sizes provide higher resolution tissue images, but require a significantly longer mass spectral acquisition time and more data storage space. The number of the image pixels is dependent on the laser raster step size to set up and the tissue size. For a typical bovine lens which has a 1 cm2 tissue size, a tissue image is typically composed of ca. 5,000 pixels if a laser raster step size of 200 &#956;m is used on an FTICR instrument. Use a &#34;random spot&#34; analysis, as this prevents location-based bias due to gradual signal attenuation during the experiment.</li>
</ol>
4. Data Analysis
<ol>
	<li>Calibrate the MALDI mass spectra using internal calibration for the initial comparison and to select peaks for MS/MS. De-isotope and select the monoisotopic peaks as previously described, using a customized VBA script38.</li>
	<li>Export the resulting monoisotopic peak lists and input the measured m/z values into the METLIN39 and/or the HMDB40 metabolome databases for mass matching with the library entries. Consider the (M+H)+, (M+Na)+, and (M+K)+ ions during the database searches, with an allowable mass error of &#177;1 ppm.</li>
	<li>Generate MALDI images for all of the lipid entities detected across the entire tissue section using image-analysis software, with a mass filter width of 1 ppm at the peak apex.</li>
	<li>Once images have been generated for all m/z values that match database entries, generate images for all other peaks as well&#160;to look for unique distribution patterns that can be investigated later.</li>
</ol>
5. Confirmation of the Identities of the Imaged Lipids
<ol>
	<li>Confirm the identities of the high abundance lipids, which have characteristic fragment ions that can be detected using the FTICR instrument (e.g.&#160;184.073 for phospholipids), by MALDI-MS/MS. Perform MALDI-MS/MS using collision-induced dissociation (CID) directly on the tissue.</li>
	<li>For those lipid species that cannot be directly confirmed by MALDI-MS/MS, use a UPLC system coupled to a q-TOF mass spectrometer34.
	<ol>
		<li>Manually dissect aliquots containing ~10 mg of tissue from the area where the species of interest was localized. Place these tissue aliquots into 2&#160;ml centrifuge tubes.</li>
		<li>Homogenize each tissue aliquot in 250 &#956;l of water, using a mixer mill with two 5 mm stainless steel metal balls.</li>
		<li>Add 1 ml of chloroform-methanol solution (1:3, v/v), and vortex the tubes. Next, centrifuge the tubes using a microcentrifuge at 12,000 x g&#160;for 10 min.</li>
		<li>Collect the supernatants and dry them in a rotary speed-vacuum concentrator.</li>
		<li>Dissolve the residues in 100 &#956;l of 30:70 isopropanol:water. Inject a 10-&#956;l aliquot onto the UPLC column for separation using gradient elution.</li>
		<li>Use the chromatographic conditions for on-line lipid LC-MS/MS, which have been published previously34.</li>
		<li>Generate extracted ion current (XIC) chromatograms using the theoretical m/z values, with a window of &#177;50 ppm around the theoretical masses.</li>
		<li>If authentic compounds for those lipids are available, match the retention times of the authentic compounds with those of the corresponding XIC peaks from the tissue samples. If the compounds are the same, the retention times and the MS/MS spectra should match.</li>
	</ol>
	</li>
	<li>If an authentic compound is unavailable, use the fragmentation pattern of the detected lipid to match a standard MS/MS spectrum&#160;from a metabolome database such as METLIN or HMDB. Use de novo mass spectral interpretation to determine a possible structure for the lipid.</li>
</ol>

We have nothing to disclose.

The most important considerations for successful MALDI MSI are: 1) tissue preparation; 2) matrix choice; 3) matrix application; and 4) data interpretation and analysis. When the sample and the matrix are appropriately prepared, the MS data acquisition is automated. The data analysis from this type of experiment is quite labor-intensive.
Appropriate tissue preparation is crucial for successful MALDI MSI experiments. The source of the tissue and the handling can have a large impact on the final ...

Mass spectrometry imaging (MSI) is an analytical technique for determining the spatial localization and distribution patterns of compounds on the surface of a tissue section1,2. Matrix assisted laser&#160;desorption/ionization (MALDI) MSI for the analysis of peptides and proteins has been used for over a decade and there have been great improvements in methods for sample preparation, detection sensitivity, spatial resolution, reproducibility and data processing3,4. By combining information from histologically stained sections and MSI experiments, pathologists are able to correlate the distributions of specific compounds with pathophysiologically interesting features5.
The distribution patterns of small molecules, including exogenous drugs6,7 and their metabolites8-10 have also been interrogated by MALDI-MS tissue imaging11. Lipids are perhaps the most widely-studied class of compounds with MALDI imaging, both in the MS12-17 and MS/MS18 modes. The use of MALDI MSI for small molecule imaging has been limited by several factors: 1) MALDI matrices are themselves small molecules (typically m/z &#60;500), which generate abundant ion signals. These abundant signals can suppress the ionization of small-molecule analytes and interfere with their detection19,20. Solvent-free matrix coating21, matrix sublimation22, and matrix precoated MALDI MS23, among others, have been developed to improve MSI of small molecules.
New matrices that can improve the analysis of low-MW compounds are of great interest in small-molecule MSI. These matrices should provide increased analyte signals with decreased matrix signals. In the positive-ion mode, 2,5-dihydroxybenzoic acid (DHB) and &#945;-cyano-4-hydroxycinnamic acid (CHCA) are the two commonly used MALDI MS matrices for MSI24. The ideal matrix would form small crystals, so as to preserve the spatial localization of the analytes. DHB tends to form larger crystals, therefore applying the matrix using sublimation has been developed to partially overcome this problem, and has allowed the use of this matrix for sensitive imaging of phospholipids22,25. 9-Aminoacridine has been used for MSI of protic analytes in the positive-ion mode26 and for nucleotides and phospholipids in the negative-ion mode26-29. 2-Mercaptobenzothiazole has been found to give efficient MALDI detection of lipids30, and has been used for the imaging of mouse&#160;brain gangliosides31. The ultrahigh resolution of Fourier transform ion cyclotron resonance (FTICR) mass spectrometers can somewhat alleviate this problem by resolving analyte signals from matrix signals32. Another advantage of the use of FTICR-MS is that the intensities of the metastable matrix clusters are reduced33, which also reduces these interferences27.
The use of dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) as a MALDI matrix for tissue imaging has previously been reported34. In this current work, a detailed protocol is provided for the use of DT for the MSI of endogenous lipids on the surfaces of bovine lens tissue sections, in the positive-ion mode.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Rat Liver</td>
			<td>Pel-Freez Biologicals</td>
			<td>56023-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Calf Lens</td>
			<td>Pel-Freez Biologicals</td>
			<td>57114-2</td>
			<td>Sample should be decapsulated29 before use</td>
		</tr>
		<tr>
			<td>Dithranol (DT)</td>
			<td>Sigma-Aldrich</td>
			<td>10608</td>
			<td>MALDI Matrix</td>
		</tr>
		<tr>
			<td>&#945;-Cyano-4-hydroxy-cinnamic Acid (CHCA)</td>
			<td>Sigma-Aldrich</td>
			<td>70990</td>
			<td>MALDI Matrix</td>
		</tr>
		<tr>
			<td>2,5-Dihydroxybenzoic Acid (DHB)</td>
			<td>Sigma-Aldrich</td>
			<td>85707</td>
			<td>MALDI Matrix</td>
		</tr>
		<tr>
			<td>Reserpine</td>
			<td>Sigma-Aldrich</td>
			<td>83580</td>
			<td></td>
		</tr>
		<tr>
			<td>Terfenadine</td>
			<td>Sigma-Aldrich</td>
			<td>T9652</td>
			<td></td>
		</tr>
		<tr>
			<td>Formic Acid</td>
			<td>Sigma-Aldrich</td>
			<td>14265</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium Formate</td>
			<td>Sigma-Aldrich</td>
			<td>14266</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium Hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>320145</td>
			<td></td>
		</tr>
		<tr>
			<td>Trifluoroacetic Acid (TFA)</td>
			<td>Sigma-Aldrich</td>
			<td>302031</td>
			<td></td>
		</tr>
		<tr>
			<td>Water</td>
			<td>Sigma-Aldrich</td>
			<td>39253</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Sigma-Aldrich</td>
			<td>34860</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetonitrile</td>
			<td>Sigma-Aldrich</td>
			<td>34967</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl Acetate</td>
			<td>Sigma-Aldrich</td>
			<td>34972</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol</td>
			<td>Sigma-Aldrich</td>
			<td>34965</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloroform</td>
			<td>Sigma-Aldrich</td>
			<td>366927</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Sigma-Aldrich</td>
			<td>34850</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Commercial Alcohols</td>
			<td></td>
			<td>95%</td>
		</tr>
		<tr>
			<td>ES Tuning Mix</td>
			<td>Agilent Technologies</td>
			<td>G2431A</td>
			<td></td>
		</tr>
		<tr>
			<td>ITO Coated Glass Slides</td>
			<td>Hudson Surface Technology</td>
			<td>PSI1207000</td>
			<td>Ensure that samples are placed on the electrically conductive side</td>
		</tr>
		<tr>
			<td>Wite-Out Shake-N-Squeeze Correction Pen</td>
			<td>Bic</td>
			<td>WOSQP11</td>
			<td></td>
		</tr>
		<tr>
			<td>Airbrush Sprayer</td>
			<td>Iwata</td>
			<td>Eclipse HP-CS</td>
			<td></td>
		</tr>
		<tr>
			<td>ImagePrep</td>
			<td>Bruker</td>
			<td>249500-LS</td>
			<td></td>
		</tr>
		<tr>
			<td>MALDI adapter</td>
			<td>Bruker</td>
			<td>235380</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

dithranol as a matrix for matrix assisted laser desorption/ionization imaging on a fourier transform ion cyclotron resonance mass spectrometer

Mass spectrometry imaging (MSI) determines the spatial localization and distribution patterns of compounds on the surface of a tissue section, mainly using MALDI (matrix&#160;assisted laser desorption/ionization)-based analytical techniques. New matrices for small-molecule MSI, which can improve the analysis of low-molecular weight (MW) compounds, are needed. These matrices should provide increased analyte signals while decreasing MALDI background signals. In addition, the use of ultrahigh-resolution instruments, such as Fourier transform ion cyclotron resonance (FTICR) mass spectrometers, has&#160;the ability to resolve analyte signals from matrix signals, and this can partially overcome many problems associated with the background originating from the MALDI matrix. The reduction in the intensities of the metastable matrix clusters by FTICR MS can also help to overcome some of the interferences associated with matrix peaks on other instruments. High-resolution instruments such as the FTICR mass spectrometers are advantageous as they can produce distribution patterns of many compounds simultaneously while still providing confidence in chemical identifications. Dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) has previously been reported as a MALDI matrix for tissue imaging. In this work, a protocol for the use of DT for MALDI imaging of endogenous lipids from the surfaces of mammalian tissue sections, by positive-ion MALDI-MS, on an ultrahigh-resolution hybrid quadrupole FTICR instrument has been provided.

Tissue samples that are sectioned and thaw mounted onto the ITO coated glass slides should be intact, without visible tearing. For many tissues, direct tissue thaw mounting onto an ITO coated glass slide is acceptable. For some specific tissues such as bovine lens, extensive tearing of the tissue is often seen when direct thaw mounting is used (Figure 1a). Precoating of the ITO glass slide with ethanol or formic acid (Figure 1b) helps to maintain the integrity of the tissue sections duri...

Watch this Scientific Journal Video about Dithranol as a Matrix for Matrix Assisted Laser Desorption/Ionization Imaging on a Fourier Transform Ion Cyclotron Resonance Mass Spectrometer at JoVE.com

Dithranol as a Matrix for Matrix Assisted Laser Desorption/Ionization Imaging on a Fourier Transform Ion Cyclotron Resonance Mass Spectrometer

Dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) has previously been reported as a MALDI matrix for tissue imaging of small molecules; protocols for the use of DT for the MALDI imaging of endogenous lipids on the surface of tissue sections&#160;by positive-ion MALDI-MS on an ultrahigh-resolution quadrupole-FTICR instrument are provided here.

Mass spectrometry imaging (MSI) determines the spatial localization and distribution patterns of compounds on the surface of a tissue section, mainly ...

dithranol-as-matrix-for-matrix-assisted-laser-desorptionionization

Research

JoVE Journal

Biology

14.1K Views.  University of Victoria.  Dithranol (DT, 1,8-dihydroxy-9 ,10-dihydroanthracen-9-on) is eerder gerapporteerd als een MALDI matrix voor weefsel beeldvorming van kleine moleculen, protocollen voor het gebruik van DT voor de MALDI beeldvorming van endogene lipiden op de oppervlak van het weefsel secties door positieve-ion MALDI-MS op een ultrahoge resolutie quadrupool-FTICR instrument worden hier gegeven. 

Video: Dithranol as a Matrix for Matrix Assisted Laser Desorption/Ionization Imaging on a Fourier Transform Ion Cyclotron Resonance Mass Spectrometer

MALDI Sample Preparation: the Ultra Thin Layer Method

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS) 1,2. The ultra-thin layer method involves the production of a substrate layer of matrix crystals (alpha-cyano-4-hydroxycinnamic acid) on the sample plate, which serves as a seeding ground for subsequent crystallization of a matrix/analyte mixture. Advantages of the ultra-thin layer method over other sample deposition approaches (e.g. dried droplet) are that it provides (i) greater tolerance to impurities such as salts and detergents, (ii) better resolution, and (iii) higher spatial uniformity. This method is especially useful for the accurate mass determination of proteins. The protocol was initially developed and optimized for the analysis of membrane proteins and used to successfully analyze ion channels, metabolite transporters, and receptors, containing between 2 and 12 transmembrane domains 2. Since the original publication, it has also shown to be equally useful for the analysis of soluble proteins. Indeed, we have used it for a large number of proteins having a wide range of properties, including those with molecular masses as high as 380 kDa 3. It is currently our method of choice for the molecular mass analysis of all proteins. The described procedure consistently produces high-quality spectra, and it is sensitive, robust, and easy to implement.

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS).

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption ...

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System 

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers. Genetic studies in mammalian systems have been limited due to the lack of readily available tools including defined mutant genetic cell lines, regulatory expression systems, and appropriate selectable markers. To circumvent these difficulties, model genetic systems in lower eukaryotes have become an attractive choice for the study of functionally conserved DNA repair proteins and pathways. We have developed a model yeast system to study the poorly defined genetic functions of the Werner syndrome helicase-nuclease (WRN) in nucleic acid metabolism. Cellular phenotypes associated with defined genetic mutant backgrounds can be investigated to clarify the cellular and molecular functions of WRN through its catalytic activities and protein interactions. The human WRN gene and associated variants, cloned into DNA plasmids for expression in yeast, can be placed under the control of a regulatory plasmid element. The expression construct can then be transformed into the appropriate yeast mutant background, and genetic function assayed by a variety of methodologies. Using this approach, we determined that WRN, like its related RecQ family members BLM and Sgs1, operates in a Top3-dependent pathway that is likely to be important for genomic stability. This is described in our recent publication [1] at <a href="http://www.impactaging.com">www.impactaging.com</a>. Detailed methods of specific assays for genetic complementation studies in yeast are provided in this paper.

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System

Genetic studies in yeast can be employed to investigate the molecular and cellular functions of human genes in cellular DNA metabolism. Methods are described for the genetic characterization of the human WRN gene product defective in the premature aging disorder Werner syndrome in functionally conserved pathways using yeast as a tractable model system.

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers.  Genetic studies in mammalian ...

Studying the Effects of Matrix Stiffness on Cellular Function using Acrylamide-based Hydrogels

Tissue stiffness is an important determinant of cellular function, and changes in tissue stiffness are commonly associated with fibrosis, cancer and cardiovascular disease1-11. Traditional cell biological approaches to studying cellular function involve culturing cells on a rigid substratum (plastic dishes or glass coverslips) which cannot account for the effect of an elastic ECM or the variations in ECM stiffness between tissues. To model in vivo tissue compliance conditions in vitro, we and others use ECM-coated hydrogels. In our laboratory, the hydrogels are based on polyacrylamide which can mimic the range of tissue compliances seen biologically12. "Reactive" cover slips are generated by incubation with NaOH followed by addition of 3-APTMS. Glutaraldehyde is used to cross-link the 3-APTMS and the polyacrylamide gel. A solution of acrylamide (AC), bis-acrylamide (Bis-AC) and ammonium persulfate is used for the polymerization of the hydrogel. N-hydroxysuccinimide (NHS) is incorporated into the AC solution to crosslink ECM protein to the hydrogel. Following polymerization of the hydrogel, the gel surface is coated with an ECM protein of choice such as fibronectin, vitronectin, collagen, etc.
The stiffness of a hydrogel can be determined by rheology or atomic force microscopy (AFM) and adjusted by varying the percentage of AC and/or bis-AC in the solution12. In this manner, substratum stiffness can be matched to the stiffness of biological tissues which can also be quantified using rheology or AFM. Cells can then be seeded on these hydrogels and cultured based upon the experimental conditions required. Imaging of the cells and their recovery for molecular analysis is straightforward. For this article, we define soft substrata as those having elastic moduli (E) &lt;3000 Pascal and stiff substrata/tissues as those with E &gt;20,000 Pascal.

The effect of substrata stiffness on cellular function can be modeled in vitro using polyacrylamide hydrogels of varying compliances.

Tissue stiffness is an important determinant of cellular function, and changes in tissue stiffness are commonly associated with fibrosis, cancer and ...

Quantifying Mixing using Magnetic Resonance Imaging

Mixing is a unit operation that combines two or more components into a homogeneous mixture. This work involves mixing two viscous liquid streams using an in-line static mixer. The mixer is a split-and-recombine design that employs shear and extensional flow to increase the interfacial contact between the components. A prototype split-and-recombine (SAR) mixer was constructed by aligning a series of thin laser-cut Poly (methyl methacrylate) (PMMA) plates held in place in a PVC pipe. Mixing in this device is illustrated in the photograph in Fig. 1. Red dye was added to a portion of the test fluid and used as the minor component being mixed into the major (undyed) component. At the inlet of the mixer, the injected layer of tracer fluid is split into two layers as it flows through the mixing section. On each subsequent mixing section, the number of horizontal layers is duplicated. Ultimately, the single stream of dye is uniformly dispersed throughout the cross section of the device.
 Using a non-Newtonian test fluid of 0.2% Carbopol and a doped tracer fluid of similar composition, mixing in the unit is visualized using magnetic resonance imaging (MRI). MRI is a very powerful experimental probe of molecular chemical and physical environment as well as sample structure on the length scales from microns to centimeters. This sensitivity has resulted in broad application of these techniques to characterize physical, chemical and/or biological properties of materials ranging from humans to foods to porous media 1, 2. The equipment and conditions used here are suitable for imaging liquids containing substantial amounts of NMR mobile 1H such as ordinary water and organic liquids including oils. Traditionally MRI has utilized super conducting magnets which are not suitable for industrial environments and not portable within a laboratory (Fig. 2). Recent advances in magnet technology have permitted the construction of large volume industrially compatible magnets suitable for imaging process flows. Here, MRI provides spatially resolved component concentrations at different axial locations during the mixing process. This work documents real-time mixing of highly viscous fluids via distributive mixing with an application to personal care products.

Magnetic resonance imaging (MRI) provides a powerful tool to evaluate the effectiveness of process equipment during operation. We discuss the use of MRI to visualize mixing in a static mixer. The application is relevant to personal care products, but can be applied to a broad range of food, chemical, biomass and biological fluids.

Mixing is a unit operation that combines two or more components into a homogeneous mixture.  This work involves mixing two viscous liquid streams using an ...

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Cellular invasion into local tissues is a process important in development and homeostasis.  Malregulated invasion and subsequent cell movement is ...

Improving the Success Rate of Protein Crystallization by Random Microseed Matrix Screening

Random microseed matrix screening (rMMS) is a protein crystallization technique in which seed crystals are added to random screens. By increasing the likelihood that crystals will grow in the metastable zone of a protein's phase diagram, extra crystallization leads are often obtained, the quality of crystals produced may be increased, and a good supply of crystals for data collection and soaking experiments is provided. Here we describe a general method for rMMS that may be applied to either sitting drop or hanging drop vapor diffusion experiments, established either by hand or using liquid handling robotics, in 96-well or 24-well tray format.

Here we describe a general method for random microseed matrix screening. This technique is shown to significantly increase the success rate of protein crystallization screening experiments, reduce the need for optimization, and provide a reliable supply of crystals for data collection and ligand-soaking experiments.

Random microseed matrix screening  (rMMS) is a protein crystallization technique in which seed crystals are added  to random screens. By increasing the ...

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of cell proliferation, survival, migration, etc. The ECM plays important roles in development and in diverse pathologies including cardio-vascular and musculo-skeletal diseases, fibrosis, and cancer. Thus, characterizing the composition of ECMs of normal and diseased tissues could lead to the identification of novel prognostic and diagnostic biomarkers and potential novel therapeutic targets. However, the very nature of ECM proteins (large in size, cross-linked and covalently bound, heavily glycosylated) has rendered biochemical analyses of ECMs challenging. To overcome this challenge, we developed a method to enrich ECMs from fresh or frozen tissues and tumors that takes advantage of the insolubility of ECM proteins. We describe here in detail the decellularization procedure that consists of sequential incubations in buffers of different pH and salt and detergent concentrations and that results in 1) the extraction of intracellular (cytosolic, nuclear, membrane and cytoskeletal) proteins and 2) the enrichment of ECM proteins. We then describe how to deglycosylate and digest ECM-enriched protein preparations into peptides for subsequent analysis by mass spectrometry.

This protocol describes a procedure for enriching ECM proteins from tissues or tumors and deglycosylating and digesting the ECM-enriched preparations into peptides to analyze their protein composition by mass spectrometry.

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of ...

Laser-assisted Microdissection (LAM) as a Tool for Transcriptional Profiling of Individual Cell Types

The understanding of developmental processes at the molecular level requires insights into transcriptional regulation, and thus the transcriptome, at the level of individual cell types. While the methods described here are generally applicable to a wide range of species and cell types, our research focuses on plant reproduction. Plant cultivation and seed production is of crucial importance for human and animal nutrition. A detailed understanding of the regulatory networks that govern the formation of the reproductive lineage (germline) and ultimately of seeds is a precondition for the targeted manipulation of plant reproduction. In particular, the engineering of apomixis (asexual reproduction through seeds) into crop plants promises great improvements, as it leads to the formation of clonal seeds that are genetically identical to the mother plant. Consequently, the cell types of the female germline are of major importance for the understanding and engineering of apomixis. However, as the corresponding cells are deeply embedded within the floral tissues, they are very difficult to access for experimental analyses, including cell-type specific transcriptomics. To overcome this limitation, sections of individual cells can be isolated by laser-assisted microdissection (LAM). While LAM in combination with transcriptional profiling allows the identification of genes and pathways active in any cell type with high specificity, establishing a suitable protocol can be challenging. Specifically, the quality of RNA obtained after LAM can be compromised, especially when small, single cells are targeted. To circumvent this problem, we have established a workflow for LAM that reproducibly results in high RNA quality that is well suitable for transcriptomics, as exemplified here by the isolation of cells of the female germline in apomictic Boechera. In this protocol, procedures are described for tissue preparation and LAM, also with regard to RNA extraction and quality control.

Laser-assisted Microdissection LAM as a Tool for Transcriptional Profiling of Individual Cell Types

Here we present a protocol for laser-assisted microdissection of specific plant cell types for transcriptional profiling. While the protocol is suitable for different species and cell types, the focus is on highly inaccessible cells of the female germline important for sexual and apomictic reproduction in the crucifer genus Boechera.

The understanding of developmental processes at the molecular level requires insights into transcriptional regulation, and thus the transcriptome, at the ...

Implementation of a Coherent Anti-Stokes Raman Scattering (CARS) System on a Ti:Sapphire and OPO Laser Based Standard Laser Scanning Microscope

Laser scanning microscopes combining a femtosecond Ti:sapphire laser and an optical parametric oscillator (OPO) to duplicate the laser line have become available for biologists. These systems are primarily designed for multi-channel two-photon fluorescence microscopy. However, without any modification, complementary non-linear optical microscopy such as second-harmonic generation (SHG) or third harmonic generation (THG) can also be performed with this set-up, allowing label-free imaging of structured molecules or aqueous medium-lipid interfaces. These techniques are well suited for in-vivo observation, but are limited in chemical specificity. Chemically selective imaging can be obtained from inherent vibration signals based on Raman scattering. Confocal Raman microscopy provides 3D spatial resolution, but it requires high average power and long acquisition time. To overcome these difficulties, recent advances in laser technology have permitted the development of nonlinear optical vibrational microscopy, in particular coherent anti-Stokes Raman scattering (CARS). CARS microscopy has therefore emerged as a powerful tool for biological and live cell imaging, by chemically mapping lipids (via C-H stretch vibration), water (via O-H stretch vibrations), proteins or DNA. In this work, we describe the implementation of the CARS technique on a standard OPO-coupled multiphoton laser scanning microscope. It is based on the in-time synchronization of the two laser lines by adjusting the length of one of the laser beam path. We present a step-by-step implementation of this technique on an existing multiphoton system. A basic background in experimental optics is helpful and the presented system does not require expensive supplementary equipment. We also illustrate CARS imaging obtained on myelin sheaths of sciatic nerve of rodent, and we show that this imaging can be performed simultaneously with other nonlinear optical imaging, such as standard two-photon fluorescence technique and second-harmonic generation.

Implementation of a Coherent Anti-Stokes Raman Scattering CARS System on a Ti:Sapphire and OPO Laser Based Standard Laser Scanning Microscope

Coherent anti-Stokes Raman scattering (CARS) microscopy based on inherent vibration of molecule bonds permits label-free chemically selective live cell imaging. This work presents the implementation of a complementary microscopy technique on a standard multiphoton laser scanning microscope based on a femtosecond Ti:sapphire laser and an OPO laser.

Laser scanning microscopes combining a femtosecond Ti:sapphire laser and an optical parametric oscillator (OPO) to duplicate the laser line have become ...

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and pathological processes. However, the isolation of ECM, from tissues or cell culture, is complicated by the insoluble and cross-linked nature of the assembled ECM and by the potential contamination of ECM extracts with cell surface and intracellular proteins. Here, we describe a method for use with cultured cells that is rapid and reliably removes cells to isolate a cell-derived ECM for downstream experimentation.
Through use of this method, the isolated ECM and its components can be visualized by in situ immunofluorescence microscopy. The dynamics of specific ECM proteins can be tracked by tracing the deposition of a tagged protein using fluorescence microscopy, both before and after the removal of cells. Alternatively, the isolated ECM can be extracted for biochemical analysis, such as sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting. At larger scales, a full proteomics analysis of the isolated ECM by mass spectrometry can be conducted. By conducting ECM isolation under sterile conditions, sterile ECM layers can be obtained for functional or phenotypic studies with any cell of interest. The method can be applied to any adherent cell type, is relatively easy to perform, and can be linked to a wide repertoire of experimental designs.

The extracellular matrix plays a major role in defining the microenvironment of cells and in modulating cell behavior and phenotype. We describe a rapid method for the isolation of cell-derived extracellular matrix, which can be adapted to different scales for microscopic, biochemical, proteomic, or functional studies.

Dithranol as a Matrix for Matrix Assisted Laser Desorption/Ionization Imaging on a Fourier Transform Ion Cyclotron Resonance Mass Spectrometer

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