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In order to perform 2-D gel analyses on restriction fragments from higher eukaryotic genomes, it is necessary to remove most of the linear, non-replicating, fragments from the starting DNA preparation. This is because the replication intermediates in a single-copy locus constitute such a minute fraction of all of the restriction fragments in a standard DNA preparation - whether isolated from synchronized or asynchronous cultures. Furthermore, the very long linear DNA strands that characterize higher eukaryotic genomes are inordinately subject to branch migration and shear. We have developed a method that results in significant enrichment of replicating fragments that largely maintain their branched intermediates. The method depends upon two important factors: 1) replicating fragments in higher eukaryotic nuclei appear to be attached to the nuclear matrix in a supercoiled fashion, and 2) partially single-stranded fragments (e.g., those containing replication forks) are selectively adsorbed to BND-cellulose in high salt concentrations. By combining matrix-enrichment and BND-cellulose chromatography, it is possible to obtain preparations that are enriched 200–300-fold over the starting genomic DNA, and are thus suitable for analysis on 2-D gels.
The neutral/neutral and neutral/alkaline two-dimensional (2-D) gel methods were originally introduced more than 20 years ago (1,2), and were utilized initially to examine the characteristics of origins of replication in S. cerevisiae. The starting material in each case was a preparation of yeast DNA from synchronized or asynchronous cultures prepared by standard CsCl banding techniques. Together, these two techniques for mapping origins and their corresponding replicons are tremendously powerful: among all the other methods for detecting origins of replication, they still afford the most detailed view of the replication intermediates inhabiting a given restriction fragment. However, the 2-D gel mapping methods were not readily applicable to replicons in the genomes of organisms more evolved than Physarum and D. melanogaster (e.g., 3,4)). This limitation derived primarily from the great complexity of higher eukaryotic genomes, the much longer cell cycle times, and the resulting very low signal-to-noise ratio of replicating to non-replicating DNA. For example, the genome of S. cerevisiae is ~300-fold less complex than the genome of a mammalian cell. Therefore, correspondingly more mammalian DNA (most of it non-replicating linear fragments) would have to be loaded into the well of a 2-D gel in order to be able to detect the intermediates in a single-copy restriction fragment. Unfortunately, it is simply not possible to effectively separate this much DNA ~1.5 mg on a 2-D gel. Furthermore, the methods routinely used to isolate and purify genomic DNA had to be modified to prevent branch migration and shear, which is a major problem with the long linear chromosomal DNA that characterizes higher eukaryotic genomes. What was needed was a method for removing the vast excess of non-replicating DNA from the fragments containing replication intermediates. With sufficient numbers of starting cells, such an enrichment step therefore would make it possible to search for origins even in mammalian genomes.
Our laboratory developed such a method, which depends upon two older observations. In the first of these, it was shown that DNA is attached at ~100 kb intervals to a proteinaceous nuclear substructure or matrix (reviewed in 5), which renders the DNA less susceptible to both branch migration and shear. The general approach is to extract nuclei with buffers containing either high salt concentrations (6) or a detergent such as lithium diiodosalicylate (LIS; 7). This treatment removes soluble nuclear proteins, histones, and most of the non-histone proteins from DNA, leaving a residual nuclear matrix to which the genomic DNA is attached. This DNA “halo” is essentially protein-free and can be digested with an appropriate restriction enzyme while still attached to the matrix.
Importantly, it also was shown that >90% of restriction fragments containing replication forks preferentially associate with the 4–5% of DNA that remains when a matrix/DNA halo preparation is digested to completion with a six-mer restriction enzyme (8,9). Therefore, by isolating the matrix-attached DNA fraction, an initial 10–20-fold enrichment of replication intermediates is obtained. A second critical observation was that partially single-stranded DNA (such as in a replication fork) is selectively adsorbed to BND-cellulose in the presence of high salt, and can subsequently be eluted with a caffeine wash (10). In practice, this second step eliminates most of the remaining linear fragments from the matrix-attached DNA fraction described above, and affords an additional 5–10-fold enrichment of RIs over linear fragments (9). Together, these two steps constitute the enrichment scheme that has allowed analyses of single-copy loci in mammalian cells by 2-D gel replicon mapping techniques on a routine basis (e.g., 11–14).
Although 2-D gel analysis of mammalian replicons has largely been supplanted by the nascent strand abundance assay, which is somewhat easier and requires less starting material owing to the sensitivity of PCR amplification (15,16), we believe that 2-D gels still afford the most comprehensive view of origin behavior. An important limitation, however, is that it is extremely difficult to detect replication bubble arcs in neutral/neutral 2-D gels in replication intermediates isolated from asynchronous cultures of mammalian cells. This is because most mammalian origins are zones of inefficient sites, and the zones themselves are inefficient. Thus, a fragment from an initiation zone will usually be replicated passively from a start site in some neighboring fragment in the zone, resulting in a strong single fork arc and a very weak bubble arc that cannot be detected on film.
Therefore, the majority of our studies have been performed on cells synchronized at the G1/S boundary, released into the S-period, and sampled at selected times thereafter. In the interest of describing the protocol from start to finish, we will detail the method of synchronizing and preparing matrices from Chinese hamster ovary (CHO) cells, which are grown in monolayers, and human lymphoid cells, which grow in suspension. These are the cells with which we have the most experience. However, the matrix enrichment method has been applied successfully to both Chinese and Syrian hamster cells (17), African Green Monkey cells (P.A. Dijkwel, unpublished), and human HeLa (L.D. Mesner, unpublished), lymphoblastoid (L.D. Mesner, unpublished), and immunoglobulin-producing, cells (14). For the latter cell types, we have not had success in arresting the population in G1 by serum or amino acid deprivation. Therefore, double thymidine blocks or a single thymidine block followed by arrest in mimosine was used to prepare cell populations arrested at the G1/S boundary (described below for lymphoblastoid cells). For other cell types, arrest in mitosis with nocodazole followed by release into medium containing mimosine might be an option. For the preparation of origin libraries by trapping bubbles in agarose, we have routinely used asynchronous cultures, but then have assessed the efficacy of the procedure by 2-D gel analysis of DNA from synchronized cells (see Chapter __, Mesner and Hamlin). We also briefly describe the modifications we have made to the neutral/neutral 2-D gel method to accommodate the larger amounts of DNA loaded onto these gels compared to experiments with yeast DNA.
With attention to the detail provided below, it will be possible for anyone familiar with the preparation of minimally-sheared DNA from mammalian cells to perfect this enrichment technique on synchronized cells. Although we describe the general method for mammalian cells, it should be applicable, in theory, to other cultured cells (e.g., insect).
The method described here is for a single time point (usually early S-phase). It would be scaled up for multiple time points or for non-synchronized cells containing fewer replication intermediates.
We thank the present and former members of our laboratory for very helpful discussions. This work was supported by a grant from the NIH to J.L.H. (RO1 GM26108).