Mass Spectrometry 793
Final Paper – Spring 2003
It is usually the case that tissue samples subjected to proteomic or genomic analysis are comprised of many different cell types. Frequently, the cells of interest are interspersed with other cells that the investigator does not want to examine. These additional cells will certainly reduce the signal of molecules detected from the cells of interest and will likely confound analysis that are focused on only some of the cells in the tissue sample. The ability to physically select cells of interest out of a tissue section is of great potential utility in many molecular biology applications and mass spectrometry is no exception. Using laser capture microdissection this is possible.
In laser capture microdissection (LCM) cells can be selected individually out of a thin slice of frozen tissue that is less than one cell (5-20 mm) thick. In LCM a thin ethylene vinyl acetate thermoplastic film is placed over the prepared tissue sample. Using a microscope, the investigator targets the cells of interest with a laser. The laser transiently heats and melts the film, adhering it to the cells below. This can be repeated to capture as few a one and as many as a few thousand cells. When the film is removed the captured cells remain adhered and are pulled out of the tissue section. When performing LCM it is necessary to prepare the sample by actively removing all water in the tissue by treating it with organic solvents such as dehydrated ethanol or xylene. Nevertheless, laser capture is a relatively gentle procedure, allowing recovery of many proteins and protein complexes, RNA transcripts several kilobases in length, and genomic DNA.
The principle limitation associated with using laser capture microdissected cells for proteomic and genomic analysis is the small sample size. Nature has provided a mechanism that allows amplification of single nucleic acid molecules, facilitating genomic analysis of microdissected samples. Proteins, on the other hand, cannot be amplified. Fortunately, mass spectrometry is an extremely sensitive detection method, and MS analysis LCM samples is currently viable. Still, the small quantities of protein analyte that are obtained via LCM are a significant limit to proteomic analysis.
In has usually been the case that proteins from tissue samples subjected to mass spectrometry are extracted into a lysis buffer. The solubilized protein is then used for ESI or MALDI MS. In addition to putting the protein in a state amenable to mass spectrometry, extraction of protein from tissue provides other benefits. The dissolved protein can be subjected to purification and separation methods that will be discussed later. Despite the benefits of extracting proteins from tissues, it does result in significant dilution. Given the small amounts of protein that are procured from LCM samples, the dilution of that protein caused by extraction or separation could prevent detection of a protein of interest.
Xu et al. describe a method by which cells obtained via LCM can be directly subjected to MALDI MS [1], obviating the need to extract proteins from the sample cells prior to mass spectrometry. After capturing cells via LCM they attach the thermoplastic film to the target analysis plate and apply sinapinic acid matrix solution directly to the sample. A portion of the cells’ protein associates with the matrix, making it amenable to ionization. The sample is then directly subjected to MALDI-TOF mass spectrometry.
This procedure has advantages to solubilizing protein prior to MS. The first advantage is that the proteins are present at greater concentrations. This translates directly to a higher signal to noise in mass spectra, which results in increased sensitivity, increased accuracy, and increased ability to detect proteins of interest. Another related advantage is that with direct MALDI a very small original sample can be used for MS. A third advantage is that direct MALDI MS of tissue samples is easier than extracting proteins prior to MS. Eliminating the extraction creates a protocol that is less time consuming, less costly, less technically challenging, and less variable.
In fact, Xu et al. are not the first group to perform direct MALDI of LCM tissue. Palmer-Troy et al. make an earlier report using a similar technique that they also use to investigate mammary carcinoma [2]. There is a difference between the two techniques with regard to the application of matrix solution to the MALDI target that Xu et al. claim gives their protocol an important advantage. Palmer-Troy et al. apply 1 ml of matrix solution to the LCM cells on the target plate. This volume far exceeds the volume of the cells. In order to restrict matrix volume, Xu et al. use a finely pulled glass capillary to deposit matrix solution on LCM cells under microscopic visualization. This allows them to employ matrix volumes of 100 pl to 10 nl. Xu et al. claim that this gives their protocol an advantage because excessive matrix volume can dilute and mix the proteins associated with each cell cluster. They also state that microspotting matrix solution provides an accurate target for the MALDI instrument’s camera system. Xu et al. do not show a direct comparison of the two matrix deposition methods, but the spectra in their paper appear to be of higher quality than those of Palmer-Troy et al.. This could be attributed to the fact that Xu et al. are using a higher performance MALDI-TOF instrument though.
In order to assess the quality of spectra obtained via their direct LCM MALDI MS protocol, Xu et al. perform comparisons between different methods of preparing the same tissue sample. Their first figure shows that histological staining prior to LCM reduces spectral quality. They show mass spectra of stained and unstained LCM captured cells. The unstained cells show much better spectral quality. This indicates that, if possible, LCM microscopy should be performed without staining.
The second figure states that the tissue dehydration process required in preparation for LCM does not result in significant protein loss from the tissues. They show spectra obtained from three samples that were unwashed, washed in 70% ethanol for one minute, and washed in water for three minutes prior to direct MALDI MS without LCM. The spectra appear similar, but this is not an adequate control. The preparation of tissue for LCM involves not only a 70% ethanol wash, but also (2 x 30 sec) 95% ethanol washes, (2 x 30 sec) 100% ethanol washes, and a (1 x 5 min) xylene wash. It is necessary to fully dehydrate the tissue in order to get good LCM results. It seems likely that the stringent dehydration washes, which they do not perform in this control, could cause protein to be lost from the tissue. Their statement, that the tissue dehydration process required in preparation for LCM does not result in significant protein loss from the tissue, is not tolerably supported.
The third figure shows the mass spectrum obtained from a LCM sample of ten cells (mouse colon crypt) prepared using their protocol. The spectrum looks good. This indicates that their protocol will perform satisfactorily on as few as ten cells.
The fourth figure shows LCM capture of unstained human breast carcinoma. The images show that almost all of the cancerous cells from the tissue section were removed without affecting the surrounding tissue. This indicates that staining tissue samples is not required for accurate LCM. This is important given their prior finding that staining reduces mass spectral quality.
The fifth figure shows the mass spectra obtained by their direct MALDI MS protocol from microdissected invasive mammary carcinoma and normal breast epithelial cells. This is a relevant comparison from a biological standpoint because epithelial cells are the primary cells from which breast cancer is derived. They show spectra over a broad m/z range from 3,000 to 70,000 units. The spectra appear generally similar with a few dozen clearly defined peaks. Though the spectra are similar, there are clear differences between them. There are a few peaks in one of the spectra that appear to be completely absent in the other. Furthermore, many peaks common to both spectra are observed at significantly different intensities. This indicates that there are differences between the protein content of these mammary carcinoma and normal epithelial samples, which is not surprising.
Comparison between different types, groups, or treatments of tissue is the basis of proteomic analysis via mass spectrometry. These analyses generally fall into two categories. The first type of analysis treats differences between the spectra being compared as markers of the samples being examined. This can facilitate categorization of unknown samples. In this case the identity and biological function of those proteins is not important. The second type of comparative proteomic analysis is concerned with the identity and biology of the changed proteins. Here the differences between the mass spectra of the samples, along with prior biological knowledge, are used as clues to help learn more about the underlying biology of the samples being examined. Xu et al. do not comment on any analysis of the differences between the cancer and normal samples. In this case the comparison between carcinoma and normal tissue provides evidence that their protocols can be used to perform comparative proteomics.
The protocol described in Xu et al. can be used for rapid whole cell proteomics, but mass spectral analysis of whole cell protein extracts has an important fundamental limitation. The mass spectra obtained from whole cell protein mixtures, even when acquired on state of the art mass spectrometers, is generally too noisy to detect most proteins of interest. The most abundant cellular proteins, and their degradation products, obscure detection of other proteins, which are present in quantities many orders of magnitude lower than the abundant proteins. Consequently, whole cell protein extracts are generally subjected to one or more separation or purification steps in combination with mass spectrometry. The reduction in complexity by separation or purification can take many different forms.
Tandem mass spectrometry is the most elegant method of separating proteins for mass spectrometry. Quadrupole magnets can be used to trap a certain bandwidth of a mass spectrum, filtering a protein mixture by m/z. In combination with CID, MSn is an excellent method to reduce mixture complexity for whole cell proteomics.
Chromatography and electrophoresis are two other methods that are commonly used to separate protein mixtures. They provide excellent results, but cause dilution of samples. Chromatography and electrophoresis can also be used to separate protein mixtures across multiple dimensions, resulting in further reduction in mixture complexity.
Affinity purification is another method that can be used to reduce the complexity of whole cell lysates. In affinity purification, a favorable chemical interaction between an immobilized substrate and the protein(s) of interest is used to pull target molecules out of a complex mixture. The form and selectivity of affinity capture methods is extremely variable. Antibodies can be used to capture a particular protein species or a group of proteins. Phosphorylated proteins can be purified using a variety of methods that convert phosphate groups to affinity handles. Nucleic acids with sequence dependant specificity can be use to capture proteins. Numerous inorganic compounds have also been demonstrated to have selective protein affinities. Affinity purification is the basis of the SELDI technique that will be discussed later.
Of these separation methods, only tandem mass spectrometry could be integrated into the direct MALDI protocol of Xu et al.. While tandem MS separates gas phase ions within the spectrometer, each of the other methods separates proteins in solution. Since the protocol of Xu et al. skips the generation of lysate, none of the solution based separation methods would be possible.
Craven et al. describe a protocol in which proteins are extracted into a lysis buffer and separated prior to mass spectrometry [3]. The protocol of Craven et al. is directly comparable to that of Xu et al. because both groups performed whole cell proteomic analysis of LCM samples, but Craven et al. used 2D-PAGE to simplify sample mixtures prior to MALDI. They captured several thousand cells from normal kidney, renal carcinoma, and cervix epithelium and extracted proteins into a urea/thiourea-based lysis buffer. They performed SDS poly-acrylamide gel electrophoresis and used silver staining to detect proteins within the gel. Each spot was identified and excised by hand, protein subjected to in-gel tryptic digest, and fragment peptides extracted. This resulted in a dilution of the whole cell protein extract into milliliters of solvent. The extract from each gel spot was analyzed via MALDI-TOF mass spectrometry and peptide fragment masses were screened against the NCBI database. Craven et al. identify between 470 and 930 gel features in their various samples, though they do not comment on how many proteins were actually identified. Additionally, Craven et al. compare the results of using several different stains for LCM imaging, but they do not sample unstained tissue. Given the evidence presented by Xu et al. that staining negatively affects spectral quality it would have been good for Craven et al. to have included unstained tissue in this control.
The 2D-PAGE protocol of Craven et al. generates far more spectral information than the direct MALDI protocol of Xu et al.. Craven et al. detect hundreds of proteins from a whole cell lysate and can obtain enough structural information to identify many of them. This compares favorably to the protocol of Xu et al., which detects only a few dozen proteins and provides very limited structural information.
While the 2D-PAGE method clearly has merits, the direct MALDI method also has strengths. The most appealing feature of the direct MALDI method is it simplicity. Assuming adequate ionization is achieved, the entire proteome could be ionized directly from tissue. Compared to the involved chemistry of 2D-SDS-PAGE and MS of tryptic digests, direct MALDI is more likely to preserve proteins in their in vivo state. Another extremely appealing characteristic of the direct MALDI method is that it allows more reasonable comparisons between proteins signals both within and across samples. In the 2D-PAGE method each protein is subject to a separate excision, digest, extraction, and MS. This makes it very difficult to control variability from these steps. In the direct MALDI method the complexity of the mass spectra serves as an internal control. Even if the direct MALDI method was naturally extended with MSn in order to detect more proteins, it would still be easier to reconstruct coherent proteome level information than it would be with 2D-PAGE and MS. It is also important to realize that Craven et al. need to obtain a few thousand cells via LCM before they have enough protein to perform 2D-PAGE. Xu et al., on the other hand, obtain quality mass spectra with as few as ten cells. There is no application of LCM that would not benefit from having to obtain 10 cells instead of 10,000. The tryptic peptide information provided by the 2D-PAGE method is currently invaluable for protein identification, but as knowledge of the proteome grows, identification of proteins without rich structural information will become easier. As bioinformatics improves, 2D-PAGE and tryptic digest will no longer be necessary to identify most proteins.
Surface enhanced laser desorption/ionization (SELDI) is another technique that allows reduction of mixture complexity for whole proteome analysis. It requires relatively small quantities of protein compared with other separation and purification techniques, which makes it attractive for analysis of LCM samples. In SELDI a complex protein mixture is applied to an affinity surface that subsequently serves as a target for MALDI. It enables selective protein retention on a wide variety of surfaces. Chromatographic surfaces include reversed phase, cationic, anionic and metal affinity while bioaffinity surfaces include antibody-antigen, DNA-protein, and receptor-ligand. Following binding of target proteins to the affinity surface, an appropriate buffer is used to wash away non-specific binding and matrix is applied. The bound proteins are then subjected to MALDI mass spectrometry. SELDI is amenable to high throughput applications and can be configured in an array format to run many assays in parallel. This makes it extremely attractive for many applications.
Batorfi et al. describe a protocol in which SELDI is used for whole cell proteome analysis from LCM tissue [4]. They capture a few thousand trophoblast cells from normal human placenta and hydatidiform mole (a malformed placental feature that generally results in termination of pregnancy) and extract whole protein into a modest 10 ml of lysis buffer. 3 ml of extract is applied to an immobilized metal affinity capture SELDI surfaces. Following washing, sinapinic acid matrix is applied to the surface and MALDI-TOF MS is performed. The IMAC surface binds proteins with exposed histidine, tryptophan, or cysteine residues. This purifies a coherent fraction of the whole cell proteome and puts the bound proteins in a state amenable to mass spectrometry. While the previous papers did not discuss the analysis of differentially detected ions, Batorfi et al. do find four peaks in their mass spectra that are significantly different between their two experimental groups. They do not comment on the biological identity or relevance of these proteins.
Of these four papers, only one makes an attempt at the analysis of the biological significance of the data that they have collected. This is an extremely important issue in current molecular biology. Many new technologies allow collection of enormous amounts of data, which must be analyzed and deciphered before they are useful. Unless technological boundaries are being pushed, the collection of proteomic and genomic data without equal investment into the analysis of that data is imprudent. Considering the problematic nature of cross-experiment comparisons, it is incumbent upon researchers to conduct experiments with data analysis in mind.
Research Proposal
Each of the methods discussed for conducting proteomic analysis of LCM tissue takes a different approach. To me, the combination of tandem MS with direct MALDI is the most exciting because of its simplicity. If I were to propose a research project for a graduate student to extend these findings, I would certainly investigate the application of LCM-MSn.
LCM of cancerous tissue is particularly appropriate for two reasons. First, cancerous tissue is a heterogeneous mix of diseased and normal tissue. Analyzing the cancer cells separately might be necessary to figure out the molecular pathology of cancer. Second, cancerous tissue is frequently available and diagnostic testing of cancer biopsies would both practical and valuable. Consequently cancer tissue is a great starting point to investigate LCM-MSn. Mammary carcinoma would be a good system because samples are available and the cell type of origin is known.
Pushing the limits of LCM-MSn would require repeated analysis of the same samples. To that end I would capture via LCM many dozen samples each of 100 cells from carcinoma and normal epithelial cells and freeze the samples at -80ºC for future use. This could be problematic because long-term storage of LCM cells may result in protein degradation. It would be important to monitor this factor and fresh LCM samples might have to be used. I would also make sure that the original tissue was fresh and handled well in order to alleviate to problem of protein degradation prior to LCM.
In order to perform tandem MS, it is necessary to define the desired bandwidth of m/z that will be captured in the first sector of the instrument. There are two approaches to this problem. One is to scan the entire range of m/z and subject each interval to a second stage of MS. The second method is to use prior knowledge to select only those bands of m/z that contain important information. I would investigate the scanning approach because it would be applicable to any type of sample.
The main problem with the scanning approach to tandem MS is that it requires collection of a separate CID mass spectrum at every bandwidth scanned. This would result in thousands of spectra and hundreds of thousands of MALDI shots for a single 2D mass spectrum. This would be difficult in terms of both sample size and time. In order to efficiently investigate high-resolution tandem MS I would limit my investigation to a small portion of the initial mass spectrum. I would choose a range of a few hundred m/z units, which contained a complex spectrum and possibly contained targets of interest that have been identified by other investigators.
The goal of this work would be to obtain a good 2D mass spectrum, over a limited range of initial m/z, using direct MALDI of LCM tissue. The separation provided by tandem MS with CID would allow resolution of far more molecular species than in the report of Xu et al.. This would be an important extension of the direct MALDI ionization protocol because separation of complex protein extracts is critical for whole cell proteomics.
References
References
1. Xu B, Caprioli R, Sanders M, Jensen R: Direct Analysis of Laser Capture Microdissected Cells by MALDI Mass Spectrometry. American Society of Mass Spectrometry 2002, 13:1292-1297
2. Palmer-Troy DE, Sarracino D, Sgroi D, LeVangie R, Leopold P: Direct Acquisition of Matrix-assisted Laser Desorption/Ionization Time-of-Flight Mass Spectra from Laser Capture Microdissected Tissues. Clinical Chemistry 2000, 46:1513-1516
3. Craven R, Totty N, Harnden P, Selby P, Banks R: Laser Capture Microdissection and Two-Dimensional Polyacrylamide Gel Electrophoresis. American Journal of Pathology 2002, 160:815-822
4. C Batorfi J, Ye B, Mok S, Cseh I, Berkowitz R, Fulop V: Protein profiling of complete mole and normal placenta using ProteinChip analysis on laser capture microdissected cells. Gynecologic Oncology 2003, 88:424-428
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