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Selective Cleaning of the Cell Debris in Human Chromosome Preparations Studied by Scanning Force Microscopy
Journal of Structural Biology 128, 200–210 (1999) Article ID jsbi.1999.4191, available online at http://www.idealibrary.com on
Selective Cleaning of the Cell Debris in Human Chromosome Preparations Studied by Scanning Force Microscopy Javier Tamayo,*,1 Mervyn Miles,* Angela Thein,† and Peter Soothill† *H. H. Wills Physics Laboratory, and †Fetal Medicine Research Unit, St. Michael’s Hospital, University of Bristol, Tyndall Avenue, Bristol BS8 1TL, United Kingdom Received June 28, 1999, and in revised form September 14, 1999
responsible for the higher order DNA compaction within the metaphase chromosome (Rattner, 1992). The chromosome structure has been a focus of study by optical microscopy (OM) and electron microscopy (EM) in the past 30 years. OM studies have provided evidence of the higher order structure of chromatin. These studies suggest that metaphase packing is achieved by helical coiling of a 200- to 300-nm fiber and that sister chromatids have opposite helical handedness (Rattner and Lin, 1985; de la Tour and Laemmli, 1988; Saitoh and Laemmli, 1994). EM has allowed high-resolution imaging of metaphase chromosomes resolving individual chromatin fibers (Harrison et al., 1983; Adolph et al., 1986). However, evidence of higher-order structure has not been provided by EM. Since the invention of the scanning force microscope (Binnig et al., 1986), many areas of science have been benefited (Miles, 1997). Thus, scanning force microscopy (SFM) is becoming an essential technique for the study of biological structures (Bustamante and Keller, 1995). There are several reasons: (i) the high spatial resolution has provided the molecular structure of several proteins and cell membranes; (ii) sample preparation, which can distort the biological structure, is not required; (iii) the capability for imaging in liquids allows the study of the native conformation of the biomolecules in physiological buffers; and (iv) several subtechniques have been recently developed, allowing the local measurement of electrostatic, mechanical, and adhesion properties (Miles, 1997; Hoh and Heinz, 1999). Thus, single molecule spectroscopy has been demonstrated (Florin et al., 1994; Lee et al., 1994). The force that binds a single antibody molecule and a single antigen molecule has been measured as the forces between complementary single DNA strands. SFM should be an ideal candidate for studying the chromosome structure and complementing the information obtained from other microscopy techniques (Fritzsche et al., 1997). Indeed, SFM has provided
The chromosome structure is one of most challenging biological structures to be discovered. Most evidence about the structure comes from optical microscopy. Scanning force microscopy (SFM) can achieve molecular resolution and allows imaging in liquids. However, little information about the chromosome structure has been revealed by SFM. In this work, a mild enzymatic treatment is applied to the chromosomes to remove selectively the RNA and proteins coming from the cell. The resulting SFM images indicate that a protein film with embedded RNA molecules covers chromosomes in standard cytogenetic preparations. The thickness of the protein layer is 15–35 nm and the RNA adheres preferentially to the chromosome surface. The cell material film results in a quite smooth chromosome surface without evidence of any structural detail. After treatment, the chromosome was cleaned from cell residues and individual chromatin fibers at the surface were resolved. Furthermore, insights about the higher order structure of the chromosome can be inferred. r 1999 Academic Press Key Words: metaphase chromosomes; chromosome structure; scanning force microscopy; chromatin. I. INTRODUCTION
One of the most intriguing and enigmatic biological structures is the chromosome. How approximately 2 m of DNA (⬃2 ⫻ 3 ⫻ 109 bp) is folded into 46 human chromosomes within a nucleus whose diameter is about 10 µm is an unanswered question. The histone–DNA and histone–histone interactions provide a 30-nm-wide basic chromatin fiber. This fiber compacts the DNA about 40-fold, but a total compaction of about 104-fold is required to reach metaphase chromosomes. There is evidence that a hierarchy of loops and coils of the chromatin fiber is 1 To whom correspondence should be addressed. E-mail: [email protected].
1047-8477/99 $35.00 Copyright r 1999 by Academic Press All rights of reproduction in any form reserved.
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3-D images of chromosomes with nanometer spatial resolution. The 3-D imaging capability has allowed chromosome classification (karyotyping) based on chromosome volume (McMaster et al., 1996; Fritzsche and Henderson, 1996). The viscoelastic properties of metaphase chromosomes in different solutions have been characterized at nanometer scale (Fritzsche and Henderson, 1997; Xu and Ikai, 1997). Furthermore, the SFM tip has proved to be a new microdissecting tool to obtain small DNA probes for in situ hybridization (ISH) (Thalhammer et al., 1997; Xu and Ikai, 1998). Surprisingly, the SFM has not provided new insights about chromosome structure. A granular surface structure with features in the range 30–100 nm has been observed (De Grooth and Putman, 1992). These structures have been tentatively associated with the chromatin fiber loops at the surface. However, the possibility of an artifact due to the cell residues of the chromosome preparation was not discounted (Fritzsche et al., 1997; De Grooth and Putman, 1992). Indeed, the standard cytogenetic preparations involve chromosomes as well as cell material. Routine harvesting leads to chromosomes embedded in an elastic protein film (Fritzsche and Henderson, 1997; Claussen et al., 1994). Furthermore, it is known that hybridization of DNA probes within chromosomes increases after RNA and protein removal; both might either interfere with hybridization reaction or cause nonspecific binding (Gall and Pardue, 1971; Hayata, 1993). In our opinion, the SFM capability to study the chromosome structure has been limited by the presence of cell material that hides the chromosome surface. In this work, an enzymatic treatment to remove the extrinsic cell material in standard cytogenetic preparations is applied to study the chromosome structure. RNase and pepsin treatments were applied to remove selectively the RNA and proteins coming from the cell, respectively. After chromosome deposition, the glass support regions between chromosomes (substrate hereafter) as well as the chromosomes are completely covered by cell material. Selective cleaning of RNA and proteins is achieved after RNase and pepsin treatments, respectively. Cell material cleaning is achieved when the pepsin is applied after the RNase treatment. The results indicate that a protein film with embedded RNA molecules covers chromosomes in standard cytogenetic preparations. Furthermore, the thickness of the protein film could be measured at about 30 nm. The chromosomes prepared by the standard cytogenetic procedure showed a quite smooth surface. However, individual chromatin fibers could be resolved after RNA removal followed by protein diges-
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tion. These grains are associated with the chromatin fiber loops at the surface. Furthermore, the chromosome showed incomplete chromatid splitting, indicating an intermediate stage between the single prophase and the double metaphase (Sumner, 1991). On other hand, some thicker/thinner regions could be identified as Giemsa positive/negative bands, especially the reverse bands at telomeres (Musio et al., 1994). These regions showed looser chromatin. These features were not visible without treatment and provide new insights into the chromatin higher order structure in late prophase and early metaphase chromosomes. II. MATERIALS AND METHODS A. Chromosome Preparation 1. Blood culture. Heparinized peripheral human blood (1 ml) was added to 17 ml of prewarmed Iscove’s modified DMEM medium with 1% GPS (L-glutamine, penicillin, and streptomycin antibiotic mix). Two milliliters of fetal calf serum and 200 µl phytohemaglutinin were also added. The solutions were gently mixed in a flask, which was placed lying flat in an incubator for 72 h at 37°C. 2. Harvesting for metaphase chromosomes (chromosome suspension preparation). One hundred microliters of colchicine was added to the culture, mixed gently, and incubated for 1 h at 37°C. The culture was transferred to a centrifuge at 1 000 rpm for 10 min. The supernatant was removed and the pellet was resuspended in leftover fluid. Five milliliters of prewarmed 0.075 M hypotonic KCl solution was gently added by pouring it down along the side wall of test tube in a dropwise manner. The pellet was thoroughly resuspended in KCl and incubated for 20 min at 37°C. The sample was centrifuged again at 1 000 rpm for 10 min. The supernatant was discarded and the pellet was resuspended in 5 ml of freshly made fixative, 3:1 methanol:acetic acid. It was centrifuged at 1 000 rpm for 10 min, and the supernatant was discarded and resuspended in fixative again. This step was repeated two to three times until the pellet and supernatant became clear. At this stage, the chromosome pellet in the fixative solution was stored at 4°C or metaphase slides were prepared as below. 3. Slide preparation. Prior to the procedure, new slides were soaked in absolute ethanol and left in a 4°C refrigerator for a few minutes. A previously prepared fixed (in fixative solution, methanol:acetic acid at 1:3) cell suspension, stored at ⫺20°C, was centrifuged at 1 500 rpm for 7 min. The supernatant was discarded and the cell pellet was resuspended in leftover fixative solution to achieve the right concentration. Slides in absolute ethanol were removed individually and dried. A drop of suspension was dropped onto the slide. The slide was air-dried. Slides were checked under a phase-contrast microscope to ensure that cell density was correct and that there were enough well-spread, cytoplasm-free mitoses. 4. RNase treatment. RNase A (Boehringer) stock solution (20 mg/ml) was diluted 1:200 in 2⫻ SSC (0.15 M NaCl, 0.015 M sodium citrate), applied to the slide (coverslip was 24 ⫻ 60 mm2 ), and incubated for 40 min at 37°C. The slide was then washed in 2⫻ SSC for 5 min three times, with shaking at room temperature. 5. Protein digestion. Ten microliters of pepsin (100 mg/ml) was added to 100 ml of 0.01 M HCl (1 ml HCl in 99 ml dH2O). Slides were incubated in pepsin solution for 5 min at 37°C. The slides were washed in PBS buffer for 10 min at room temperature followed by a single wash in PBS/MgCl2 for 5 min. Finally, the
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slides were dehydrated in an alcohol series prior to analysis under a scanning force microscope. B. Scanning Force Microscopy A commercial scanning force microscope was used (D3100, Digital Instruments, Santa Barbara, CA, www.di.com). This scanning force microscope is equipped with an optical microscope and a motorized stage with micrometer precision to move the sample with respect to the tip. Both allow the precise location of the chromosome spreads on the slide. The measurements were taken in air with a relative humidity of 40–50% during the experiments. The scanning force microscope was operated in contact mode. No significance improvements were observed in the spatial resolution by using dynamic modes such as intermittent contact mode or noncontact mode. It should be noted that the chromosomes in air are dehydrated and are hard enough to be imaged in contact mode without any damage (Fritzsche and Henderson, 1997). V-shaped silicon nitride cantilevers were used with the lowest accessible normal spring constant, 0.06 N/m (nominal value). The tips were oxide-sharpened. The normal forces between the tip and the sample were kept below 20 nN. During the topography data acquisition, the frictional force between the tip and the sample was also measured. Differences in the frictional force were associated with compositional variations of the surface (Overney et al., 1992; Tamayo et al., 1996). III. RESULTS
A. Effect of the Treatment on the Substrate To evaluate the effect of the enzymes on the chromosome structure, topographic images of the regions between chromosomes (substrate) were taken. The flatness of the substrate allows the determination of the effect of both enzymes, RNase and pepsin, in the cell material adsorbed on the glass support. Figures 1a and 1b show the images of the glass support after chromosome deposition in two different preparations. Though the cell material adsorption on glass is different in both preparations, there are common features. The surface has a significant roughness of 2.5–2.7 nm in comparison to the glass (⬃0.3 nm). This indicates that cell material has been adsorbed on the glass support after chromosome deposition. The rough surface consists of diffuse protuberances with a characteristic size of about 200 nm. On the other hand, holes can be seen on the substrate surface. The holes are larger in Fig. 1a with a diameter of 100–150 nm and a depth of about 25 nm. The hole diameter and depth in Fig. 1b are about 50 and 20 nm, respectively. The effect of the RNA removal from the substrate is shown in Fig. 1c. The substrate showed a granular structure with a grain size of about 50 nm with the same holes as before RNase treatment. The change in the surface is a result of the removal of the diffuse protuberances of the untreated substrate. This suggests that these protuberances can be associated with RNA molecules. In fact, when protein digestion by pepsin is applied to the untreated substrate, the diffuse protuberances appear as well-defined is-
lands, indicating that these islands are composed mainly of RNA (Fig. 1d). Figure 1e shows the effect of protein digestion on the RNase-treated substrate shown in Fig. 1c. A glass support free of cell material is achieved. The glass identification is based on its well-known slight roughness. Some residual particles from the treatments appear adsorbed on the glass. The holes seen before protein digestion (Fig. 1c) disappear. If the holes in the substrate after RNA digestion go through the protein layer to the glass, the depth of these holes would be the protein film thickness. To examine this possibility, friction force experiments were performed in larger holes in the substrate after RNase treatment (data not shown). The frictional force between the tip and the sample was different on the bottom of some holes and the substrate, indicating different compositions (Overney et al., 1992; Tamayo et al., 1996). This suggests that the bottom of these holes is the bare glass and indicates that the protein layer thickness in this preparation is 29–32 nm. When protein digestion by pepsin is applied after chromosome deposition, two effects are visible on the substrate (Fig. 1d). First, the holes disappear. This is consistent with the removal of a protein layer covering the sample as was described before. The other effect is that islands in the same position as the diffuse protuberances in the untreated substrate cover partially the glass substrate. The islands are clusters of globular-like units with a diameter of 50–60 nm. The island height is 12–20 nm. Figure 1f shows the effect of RNase treatment on the pepsin-treated surface shown in Fig. 1d. The islands that covered the glass before RNA digestion surprisingly remain. However, the size of the islands decreased. The diameter of island subunits and the island height were reduced to 40–50 and 7–12 nm, respectively. Although protein digestion and RNA digestion were applied in both preparations (Figs. 1a and 1b), the order of the treatments has an influence on the removal of the cell material adsorbed after chromosome deposition. RNA removal followed by protein digestion removes most of the cell material adsorbed on the glass support (Fig. 1e). However, cell material removal remains incomplete when RNase treatment is applied after pepsin treatment. The origin of this effect will be discussed under Section IV. B. Effect of the Treatment on the Chromosome Structure Up to now, the effect of the RNA and protein digestions in the regions between chromosomes in standard cytogenetic preparations has been analyzed. The adsorbed cell material on the glass is
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FIG. 1. (a and b) A region between chromosomes for two different standard cytogenetic preparations. (c and e) The region in a after RNase treatment and following pepsin treatment, respectively. (d and f) The region in b after pepsin treatment and following RNase treatment, respectively. The Z range is 50 nm. Scan size is 3.6 ⫻ 3.6 µm2.
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removed after, first, RNase treatment and, second, pepsin treatment. The consequences of this treatment on the chromosome morphology are illustrated in Fig. 2, which shows the image of an untreated chromosome (Figs. 1a and 1b), after the RNase treatment (Figs. 1c and 1d), and after the subsequent pepsin treatment (Figs. 1e and 1f). The lefthand images show the overall chromosome structure and the right-hand images show higher resolution images of the chromosome surface. Figure 2a shows the characteristic chromosome topography in standard cytogenetic preparations. The average chromosome width and height are 1.4–1.5 µm and 98 nm, respectively. These values are in agreement with other SFM studies (Fritzsche et al., 1997). The higher resolution image shows a quite smooth surface without evidence of chromatin fibers (Fig. 2b). The effect of RNA removal is shown in Figs. 2c and 2d. The average chromosome height decreased to 70 nm. A change from a smooth to a rough surface is produced (Fig. 2d). Both results indicate that a significant amount of RNA material was covering the chromosome. This can be seen in the changes in the substrate surface after RNA digestion (compare Figs. 1a and 1c). The chromosome surface after RNase and subsequent pepsin treatments is shown in Figs. 2e and 2f. The chromosome shows a well-defined centromere and chromatids (Fig. 2e). The chromatids are not completely separated. In some regions the chromatids are bridged and in others appear almost separated. The chromosome width is unchanged after the treatment, 1.5 µm. This value is the limiting width that separates prophase chromosomes that have not split into chromatids from metaphase chromosomes with separated chromatids based on electron microscopy studies (Sumner, 1991). Therefore, this chromosome is in an intermediate state between late prophase and early metaphase in which the chromatids are separating. Variations in the average chromosome thickness of about 5–15 nm along the chromatids are also seen (Fig. 3a). Musio et al. (1994) observed that standard chromosomes showed a height pattern similar to the G-banding pattern; i.e., thicker chromosome regions correspond to G bands. The chromosome length and the centromere position suggest that the images shown in Fig. 2 correspond to chromosome 8. Indeed, some optical G bands of chromosome 8 can be identified with the thicker chromosome regions. However, several G and R bands are difficult to distinguish using the height criteria. The most evident R bands are located at telomeres. The telomere regions are clearly thinner than the rest of chromosome, about 15 nm lower than the identified G bands. Furthermore, the images suggest a telomere region
structure different from the rest of the chromosome. This structure is defined by a dense nucleus surrounded by looser chromatin (Fig. 3b). At high resolution, the chromosome surface shows a granular structure with a grain size of 45–55 nm (Figs. 2f and 3b). This size is in agreement with the diameter of the chromatin fiber, 30 nm. Furthermore, the chromosome surface is clear of cellular material that could lead to mistaken conclusions (see Fig. 1e). It is therefore reasonable to relate each grain to the chromatin fiber at the surface folding back into the chromatid body (Harrison et al., 1983; Adolph et al., 1986). The tip-sample convolution explains the larger size of the grains compared to the chromatin fiber diameter. In SFM, the horizontal dimensions of objects whose size is comparable to the tip’s radius (5–50 nm) are overestimated (Keller and Franke, 1993). The changes in the chromosome structure when the protein digestion is applied before RNase treatment are shown in Fig. 4. The chromosomes prepared by the standard cytogenetic procedure show a width and height of 1.4–1.5 µm and 115–120 nm, respectively (Fig. 4a). No evidence of chromatin fibers was observed. After protein digestion by pepsin, a rougher chromosome surface appears (Fig. 4b). The chromosome width and height remained approximately the same. Figure 4a shows the chromosome morphology after pepsin treatment followed by RNase treatment. The chromosome width increased to 1.7 µm and the height decreased to approximately 105 nm. In this case, no splitting of the chromosome into chromatids is seen. This is consistent with a late prophase stage of the chromosome. At high resolution, a well-defined granular structure was not observed as in the chromosome shown in Figs. 2e and 2f. In fact, the cell material digestion was incomplete when the RNase treatment was performed after the pepsin treatment. Substrate images showed nanometer-size islands partially covering the glass support (compare Figs. 1b, 1d, and 1f). IV. DISCUSSION
A. RNA and Protein Digestion The analysis of the changes in the substrate after each enzymatic treatment reveals that a protein layer with embedded RNA molecules covers the chromosomes in standard cytogenetic preparations. The RNA molecules appear as diffuse protuberances on the untreated substrate (Figs. 1a and 1b). The diffuse aspect indicates that the RNA is embedded in the protein film. There are two reasons to associate these protuberances with RNA molecules: (i) After RNA digestion, the diffuse protuberances disap-
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FIG. 2. Topographic images of the same metaphase chromosome after each treatment step at different magnifications. (a and b) The untreated chromosome. (c and d) The RNase-treated chromosome. (e and f) The RNase- and subsequent pepsin-treated chromosome. (a, c, and e) Scan size, 10 ⫻ 5 µm2. (b, d, and f) Scan size, 4 ⫻ 4 µm2. The force was kept below 20 nN. The Z range is 150 nm. The height offset is set to highlight the surface features hiding the lower regions of the chromosome surface. Therefore the chromosome height cannot be deduced from the images. The values are specified in the text.
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FIG. 3. Images of the chromosome shown in Figs. 2e (a) and 2f (b). This chromosome has been treated with RNase and subsequently with pepsin. In both images, the grayscale was inverted and the contrast was increased to reflect the changes in the chromosome thickness. The size and the centromere position indicate that it is chromosome 8. Scale bar is 1 µm.
peared (Fig. 1c). (ii) After protein digestion, the diffuse islands appear well defined and adsorbed on the glass (Fig. 1d). However, the islands covering the glass after pepsin treatment were not removed after RNA digestion, although their size was reduced (Fig.
1f). This suggests that some proteins are somehow protected by RNA during the protein digestion. Tentatively, these proteins could belong to polysomes (Christensen, 1994). In fact, the dimensions of the globular units that compose the islands after protein
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digestion are in agreement with the ribosome size determined by SFM (Fritzsche and Henderson, 1998). A change from a smooth chromosome surface to a rough surface is produced after RNA removal. This indicates that a significant amount of RNA covers the chromosomes. In fact, a chromosome height reduction of about 20 nm is produced after RNA removal (Figs. 2a and 2c). A decrease in chromosome height of 10–30 nm has been found in other experiments. These results suggest that clusters of RNA molecules with a thickness greater than 20 nm cover the chromosomes in standard cytogenetic preparations. This is consistent with the tendency of the ribosomes to aggregate to the chromosome surface in solution (Marsden and Laemmli, 1979). Holes can often be seen on the substrate in standard cytogenetic preparations. These holes remain after RNA digestion. Some of these holes are connected to the glass support, allowing the estimation of the thickness of the protein film in standard cytogenetic preparations. Thus, the untreated chromosome shown in Fig. 2a is covered by a protein layer with a thickness of approximately 30 nm. Measurements in holes of other samples indicate that the standard cytogenetic procedure involves chromosomes embedded in a protein layer with a thickness of 15–35 nm. B. Chromosome Structure
FIG. 4. Topographic images of a chromosome without treatment (a), after pepsin treatment (b), and after subsequent RNase treatment (c). Scan size, 7 ⫻ 7 µm2.
In standard cytogenetic preparations, the chromosome surface did not show evidence of chromatin fibers. However, after RNA and subsequent protein digestion a granular structure is observed on the chromosome surface (Figs. 2f and 3b). We have two reasons to associate each grain with the chromatin fiber folding back into the chromatid. First, the size of the grains is about 50 nm, in agreement with the chromatin fiber size. Second, the chromosome has been cleaned of cell residues, mainly RNA and proteins. Indeed, the experiments discussed in the last section indicate that a protein and RNA film with a thickness greater than 50 nm covers the untreated chromosome shown in Fig. 2a. Thus, the chromatin fibers were hidden by the cell material film, resulting in a relatively smooth chromosome surface. Other SFM studies have claimed to image grains in the chromosome surface with a size similar to that of the chromatin fiber (De Grooth and Putman, 1992). However, this granular surface has not been reproducible. Furthermore, these features were always suspected to be artifacts due to salt and cellular debris (Fritzsche et al., 1997, De Grooth and Putman, 1992). In fact, we have found some untreated chromosomes with a granular structure with a grain size of 50–100 nm. The grains could be tentatively associated with the chromatin fiber cov-
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ered by a very thin protein and RNA layer. However, the substrate showed a similar grain structure. Indeed, the protein film that covers the glass regions between chromosomes showed a granular structure with a grain size of about 50 nm (Fig. 1c). Furthermore, in scanning electron microscopy studies, a brief trypsin treatment was required to image individual chromatin fibers in standard cytogenetic preparations (Harrison et al., 1983, Sumner, 1991). A very rich chromosome structure is achieved after RNA and protein digestion. It has been stated that prophase chromosomes are longitudinally split in two chromatids. However, in 1991, Sumner showed images of late prophase chromosome without a split into chromatids. Here, high-resolution images of a chromosome in an intermediate stage between singlechromatid prophase chromosomes and a doublechromatid metaphase stage have been shown. A pattern of transverse chromatin bands bridging both chromatids can be seen in the topographic images. The study of the chromatid splitting process is significant in the elucidation of the condensation process of metaphase chromosomes. Further work will be undertaken to study the chromosome structure in different prophase and metaphase stages. A chromosome height variation of 5–10 nm along its long axis is observed (Fig. 3a). Several thicker regions correspond to G and Q bands, consistent with previous SFM observations (Musio et al., 1994). However, the correspondence between optical bands and chromosome structure does not always relate to the chromosome thickness criteria. G bands seem to fulfill the criteria of thicker chromosome regions and joined chromatids. Further work is in progress to relate the optical banding and the chromosome structure. The most evident R bands are at telomeres. These bands are thinner than the rest of the chromosome. Indeed, the long-arm telomere region is 17 nm thinner than the adjacent G band. However, the chromosome thickness variation in the internal positions is less than 10 nm. Also, a looser chromatin conformation can be inferred in the telomeres. This is consistent with the higher transcriptional activity of the R bands (Holmquist, 1992). C. Structure Nativeness A protein film with embedded RNA molecules covers chromosomes prepared by the standard cytogenetic preparation. The RNA digestion followed by protein digestion by pepsin removes most of the cell material covering the chromosome. However, this treatment can alter the chromosome composition and structure. The chromosome consists of DNA as well as proteins. The chromosomal proteins can be divided into histones (H1, H2A, H2B, H3, and H4) and nonhistone proteins. Histones are responsible for the assembly of DNA into a 30-nm chromatin
fiber. Nonhistone proteins participate in the higherorder organization of the chromatin fiber and are responsible for the overall chromosome morphology (Adolph et al., 1977; Paulson and Laemmli, 1977). Untreated chromosomes lose mainly part of histone H1 after ethanol/acetic acid fixation (Burkholder and Duczek, 1982). However, this does not seem to produce any chromatin fiber disruption (Harrison et al., 1983). The protein digestion by pepsin is the most critical step. Although protein digestion by pepsin is considered a mild treatment, we have not found biochemical data on its activity. The pepsin treatment produces the digestion of a protein layer covering the sample with a thickness of 15–35 nm. It is also reasonable to consider that some chromosomal proteins can be removed or altered. However, the chromosome showed a granular structure identified with chromatin fiber loops at the surface after RNase treatment and subsequent pepsin treatment. This structure is similar to the chromosome structure without protein extraction studied by transmission electron microscopy (TEM) (Adolph et al., 1986). Furthermore, TEM studies of histone-depleted chromosomes in solution showed a scaffolding structure surrounded by DNA loops with a diameter of 10–30 µm (Paulson and Laemmli, 1977). Clearly, these loops are not observed in this work. These observations indicate that histones in chromatin fibers are hardly affected by the treatment. Nonhistone proteins are more resistant to protease treatments than the histones (Adolph et al., 1977). The chromosomes after RNA and subsequent protein digestions showed a compact morphology. Imaging in solution also showed a compact structure (Tamayo and Miles, in press). This indicates that the main nonhistone proteins remain after the treatment. In more detail, the beginning of nonhistone removal produces a filamentous morphology to the chromatin due to separation and unfolding of the loops of the chromatin (Harrison et al., 1983). This kind of structure was not observed here, indicating that the nonhistone proteins responsible for the chromosome morphology remain after the treatment. Similar chromosome structures were obtained with a threefold pepsin concentration and double the incubation time. However, the used pepsin concentration and an incubation time slightly longer than 3 min are enough to remove the protein film. The RNA removal does not affect the chromosome structure (Adolph et al., 1977). However, the SSC solution in which the RNase is diluted extracts some nonhistone proteins (Burkholder and Duczek, 1982). Indeed, when the RNase treatment is applied after protein digestion, the chromosome width increases slightly, from 1.5 to 1.7 µm. This suggests some chromatin loosening due to the extraction of some
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nonhistone proteins. However, an increase in chromosome width was not observed when the RNase was applied to the untreated chromosome, suggesting that the protein layer covering the chromosomes screens the effect of SSC. Then, the RNA removal step does not affect the chromosomal proteins when it is applied before protein digestion. V. CONCLUSIONS
The SFM provides topographic images with high spatial resolution. Horizontal and vertical resolutions of 1 and 0.01 nm are currently obtained, respectively. However, no new information about the chromosome structure has been obtained by SFM. Furthermore, most of the known information about the chromatin structure comes from optical microscopy. The optical spatial resolution is two orders of magnitude lower than that with SFM. However, it is sensitive to the chromosome subsurface avoiding the surface contaminants. In our opinion, removal of cell material is a prerequisite to resolve the chromatin structure by SFM. RNA removal and subsequent protein digestion by pepsin treatment provided the following results: (I) Standard cytogenetic preparations consist of chromosomes covered by a protein layer with a thickness of 15–35 nm. RNA molecules are embedded in this protein layer and adhere preferentially to the chromosome surface. (II) The glass support regions between chromosomes were free of cell material after treatment. A well-defined chromosome structure was obtained and the chromatin fiber loops were resolved. Individual chromatin fibers could not be resolved in the untreated chromosomes. (III) The chromosome structure between late prophase and early metaphase is shown. The chromosome was not split into chromatids. A pattern of bridges joining the chromatids was observed. (IV) The telomeres showed a differential structure. They were clearly thinner than the rest of the chromosome and showed a looser chromatin conformation. This work was supported by the European Union TMR program. REFERENCES Adolph, K. W., Cheng, S. M., and Laemmli, U. K. (1977) Role of nonhistone proteins in metaphase chromosome structure, Cell 12, 805–816. Adolph, K. W., Kreisman, L. R., and Kuehn, R. L. (1986) Assembly of chromatin fibers into metaphase chromosomes analyzed by transmission electron microscopy and scanning electron microscopy, Biophys. J. 49, 221–231. Binnig, G., Quate, C. F., and Gerber, Ch. (1986) Atomic force microscopy, Phys. Rev. Lett. 56, 930–933.
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Burkholder, G. D., and Duczek, L. L. (1982) The effect of chromosome banding techniques on the proteins of isolated chromosomes, Chromosoma 87, 425–435. Bustamante, C., and Keller, D. (1995) Scanning force microscopy in biology, Phys. Today 32, 32–38. Christensen, A. K. (1994) Negatively-stained polysomes on rough microsome vesicles viewed by electron microscopy: Further evidence regarding the orientation of attached ribosomes, Cell Tissue Res. 276, 439–444. Claussen, U., Mazur, A., and Rubtsov, N. (1994) Chromosomes are highly elastic and can be stretched, Cytogenet. Cell Genet. 66, 120–125. De Grooth, B. G., and Putman, C. A. (1992) High-resolution imaging of chromosome-related structures by atomic force microscopy, J. Microsc. 168, 239–247. de la Tour, E. B., and Laemmli, U. K. (1988) The metapahase scaffold is helically folded: Sister chromatids have predominantly opposite helical handedness, Cell 55, 937–944. Florin, E. L., Moy, M. T., and Gaub, H. E. (1994) Adhesion forces between individual ligand–receptor pairs, Science 264, 415– 417. Fritzsche, W., and Henderson, E. (1996) Volume determination of human metaphase chromosomes by scanning force microscopy, Scanning Microsc. 10, 103–110. Fritzsche, W., and Henderson, E. (1997) Mapping elasticity of rehydrated chromosomes by scanning force microscopy, Ultramicroscopy 69, 191–200. Fritzsche, W., and Henderson, E. (1998) Ribosomes substructure investigated by scanning force microscopy and image processing, J. Microsc. 189, 50–56. Fritzsche, W., Takac, L., and Henderson, E. (1997) Application of atomic force microscopy to visualization of DNA, chromatin and chromosomes, Crit. Rev. Eukaryotic Gene Express. 7, 231–240. Gall, J. G., and Pardue, M. L. (1971) in Grosmann, L., and Moldave, L. (Eds.), Methods in Enzymology, Part D, Nucleic Acids, Vol. 21, pp. 470–480, Academic Press, New York. Harrison, C. J., Allen, T. D., and Harris, R. (1983) Scanning electron microscopy of variations in human metaphase chromosome structure revealed by Giemsa banding, Cytogenet. Cell Genet. 35, 21–27. Hayata, I. (1993) Removal of stainable cytoplasmic substances from cytogenetic slide preparations, Biotechnique Histochem. 68, 150–152. Hoh, J. H., and Heinz, W. F. (1999) Spatially resolved force spectroscopy of biological surfaces using the atomic force microscope, TIBECTH 17, 143–150. Holmquist, G. P. (1992) Chromosome bands, their chromatin flavors and their functional features, Am. J. Hum. Genet. 51, 17–37. Keller, D. J., and Franke, F. S. (1993) Envelope reconstruction of probe microscope images, Surf. Sci. 294, 409–419. Lee, G. U., Chrisey, L. A., and Colton, R. J. (1994) Direct measurement of the forces between complementary strands of DNA, Science 266, 771–773. Marsden, M. P. F., and Laemmli, U. K. (1979) Metaphase chromosome structure: Evidence of radial loop model, Cell 17, 849–858. McMaster, T. J., Winfield, M. O., Baker, A. A., Karp, A., and Miles, M. J. (1996) Chromosome classification by atomic force microscopy volume measurement, J. Vac. Sci. Technol. B 14, 1438– 1443. Miles, M. (1997) Probing the future, Science 277, 1845–1846. Musio, A., Mariani, T., Frediani, C., Sbrana, I., and Ascoli, C. (1994) Longitudinal patterns similar to G-banding in mammalian metaphase chromosomes, Chromosoma 103, 225–229.
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Overney, R. M., Meyer, E., Frommer, J., Brodbeck, D., Lu¨thi, R., Howald, L., Gu¨ntherodt, H.-J., Fujihira, M., Takano, H., and Goth, Y. (1992) Friction measurements on phase-separated thin films with a modified atomic force microscopy, Nature 359, 133–134. Paulson, J. R., and Laemmli, U. K. (1977) The structure of histone-depleted metaphase chromosomes, Cell 12, 817–828. Rattner, J. B. (1992) Integrating chromosome structure with function, Chromosoma 101, 259–264. Rattner, J. B., and Lin, C. C. (1985) Radial loops and helical coils coexist in metaphase chromosomes, Cell 42, 291–296. Saitoh, Y., and Laemmli, U. K. (1994) Metaphase chromosome structure: Bands arise from a differential folding path of the highly AT-rich scaffold, Cell 76, 609–622. Squarzoni, S., Cinti, C., Santi, S., Valmori, A., and Maraldi, N. M. (1994) Preparation of chromosome spreads for electron (TEM, SEM, STEM), light and confocal microscopy, Chromosoma 103, 381–392. Sumner, A. T. (1991) Scanning electron microscopy of mammalian chromosomes from prophase to telophase, Chromosoma 100, 410–418.
Sumner, A. T., and Ross, A. (1989) Factors affecting preparation of chromosomes for scanning electron microscopy, Scanning Microsc. Suppl. 3, 87–99. Tamayo, J., Gonza´lez, L., Gonza´lez, Y., and Garcı´a, R. (1996) Compositional mapping of semiconductor heterostructures by friction force microscopy, Appl. Phys. Lett. 68, 340–342. Tamayo, J., and Miles, M. Ultramicroscopy, in press. Thalhammer, S., Stark, R. W., Mu¨ller, S., Wienberg, J., and Heckl, W. M. (1997) The atomic force microscope as a new microdissecting tool for the generation of genetic probes, J. Struct. Biol. 119, 232–237. Xu, X.-M., and Ikai, A. (1997) Measurement of mechanical properties of human chromosomes using atomic force microscopy, Cell Eng. 2, 75–81. Xu, X.-M., and Ikai, A. (1998) Retrieval and amplification of single-copy genomic DNA from nanometer region of chromosomes: A new potential application of atomic force microscopy in genomic research, Biochem. Biophys. Res. Commun. 248, 744– 748.
Selective Cleaning of the Cell Debris in Human Chromosome Preparations Studied by Scanning Force Microscopy Javier Tamayo,*,1 Mervyn Miles,* Angela Thein,† and Peter Soothill† *H. H. Wills Physics Laboratory, and †Fetal Medicine Research Unit, St. Michael’s Hospital, University of Bristol, Tyndall Avenue, Bristol BS8 1TL, United Kingdom Received June 28, 1999, and in revised form September 14, 1999
responsible for the higher order DNA compaction within the metaphase chromosome (Rattner, 1992). The chromosome structure has been a focus of study by optical microscopy (OM) and electron microscopy (EM) in the past 30 years. OM studies have provided evidence of the higher order structure of chromatin. These studies suggest that metaphase packing is achieved by helical coiling of a 200- to 300-nm fiber and that sister chromatids have opposite helical handedness (Rattner and Lin, 1985; de la Tour and Laemmli, 1988; Saitoh and Laemmli, 1994). EM has allowed high-resolution imaging of metaphase chromosomes resolving individual chromatin fibers (Harrison et al., 1983; Adolph et al., 1986). However, evidence of higher-order structure has not been provided by EM. Since the invention of the scanning force microscope (Binnig et al., 1986), many areas of science have been benefited (Miles, 1997). Thus, scanning force microscopy (SFM) is becoming an essential technique for the study of biological structures (Bustamante and Keller, 1995). There are several reasons: (i) the high spatial resolution has provided the molecular structure of several proteins and cell membranes; (ii) sample preparation, which can distort the biological structure, is not required; (iii) the capability for imaging in liquids allows the study of the native conformation of the biomolecules in physiological buffers; and (iv) several subtechniques have been recently developed, allowing the local measurement of electrostatic, mechanical, and adhesion properties (Miles, 1997; Hoh and Heinz, 1999). Thus, single molecule spectroscopy has been demonstrated (Florin et al., 1994; Lee et al., 1994). The force that binds a single antibody molecule and a single antigen molecule has been measured as the forces between complementary single DNA strands. SFM should be an ideal candidate for studying the chromosome structure and complementing the information obtained from other microscopy techniques (Fritzsche et al., 1997). Indeed, SFM has provided
The chromosome structure is one of most challenging biological structures to be discovered. Most evidence about the structure comes from optical microscopy. Scanning force microscopy (SFM) can achieve molecular resolution and allows imaging in liquids. However, little information about the chromosome structure has been revealed by SFM. In this work, a mild enzymatic treatment is applied to the chromosomes to remove selectively the RNA and proteins coming from the cell. The resulting SFM images indicate that a protein film with embedded RNA molecules covers chromosomes in standard cytogenetic preparations. The thickness of the protein layer is 15–35 nm and the RNA adheres preferentially to the chromosome surface. The cell material film results in a quite smooth chromosome surface without evidence of any structural detail. After treatment, the chromosome was cleaned from cell residues and individual chromatin fibers at the surface were resolved. Furthermore, insights about the higher order structure of the chromosome can be inferred. r 1999 Academic Press Key Words: metaphase chromosomes; chromosome structure; scanning force microscopy; chromatin. I. INTRODUCTION
One of the most intriguing and enigmatic biological structures is the chromosome. How approximately 2 m of DNA (⬃2 ⫻ 3 ⫻ 109 bp) is folded into 46 human chromosomes within a nucleus whose diameter is about 10 µm is an unanswered question. The histone–DNA and histone–histone interactions provide a 30-nm-wide basic chromatin fiber. This fiber compacts the DNA about 40-fold, but a total compaction of about 104-fold is required to reach metaphase chromosomes. There is evidence that a hierarchy of loops and coils of the chromatin fiber is 1 To whom correspondence should be addressed. E-mail: [email protected].
1047-8477/99 $35.00 Copyright r 1999 by Academic Press All rights of reproduction in any form reserved.
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3-D images of chromosomes with nanometer spatial resolution. The 3-D imaging capability has allowed chromosome classification (karyotyping) based on chromosome volume (McMaster et al., 1996; Fritzsche and Henderson, 1996). The viscoelastic properties of metaphase chromosomes in different solutions have been characterized at nanometer scale (Fritzsche and Henderson, 1997; Xu and Ikai, 1997). Furthermore, the SFM tip has proved to be a new microdissecting tool to obtain small DNA probes for in situ hybridization (ISH) (Thalhammer et al., 1997; Xu and Ikai, 1998). Surprisingly, the SFM has not provided new insights about chromosome structure. A granular surface structure with features in the range 30–100 nm has been observed (De Grooth and Putman, 1992). These structures have been tentatively associated with the chromatin fiber loops at the surface. However, the possibility of an artifact due to the cell residues of the chromosome preparation was not discounted (Fritzsche et al., 1997; De Grooth and Putman, 1992). Indeed, the standard cytogenetic preparations involve chromosomes as well as cell material. Routine harvesting leads to chromosomes embedded in an elastic protein film (Fritzsche and Henderson, 1997; Claussen et al., 1994). Furthermore, it is known that hybridization of DNA probes within chromosomes increases after RNA and protein removal; both might either interfere with hybridization reaction or cause nonspecific binding (Gall and Pardue, 1971; Hayata, 1993). In our opinion, the SFM capability to study the chromosome structure has been limited by the presence of cell material that hides the chromosome surface. In this work, an enzymatic treatment to remove the extrinsic cell material in standard cytogenetic preparations is applied to study the chromosome structure. RNase and pepsin treatments were applied to remove selectively the RNA and proteins coming from the cell, respectively. After chromosome deposition, the glass support regions between chromosomes (substrate hereafter) as well as the chromosomes are completely covered by cell material. Selective cleaning of RNA and proteins is achieved after RNase and pepsin treatments, respectively. Cell material cleaning is achieved when the pepsin is applied after the RNase treatment. The results indicate that a protein film with embedded RNA molecules covers chromosomes in standard cytogenetic preparations. Furthermore, the thickness of the protein film could be measured at about 30 nm. The chromosomes prepared by the standard cytogenetic procedure showed a quite smooth surface. However, individual chromatin fibers could be resolved after RNA removal followed by protein diges-
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tion. These grains are associated with the chromatin fiber loops at the surface. Furthermore, the chromosome showed incomplete chromatid splitting, indicating an intermediate stage between the single prophase and the double metaphase (Sumner, 1991). On other hand, some thicker/thinner regions could be identified as Giemsa positive/negative bands, especially the reverse bands at telomeres (Musio et al., 1994). These regions showed looser chromatin. These features were not visible without treatment and provide new insights into the chromatin higher order structure in late prophase and early metaphase chromosomes. II. MATERIALS AND METHODS A. Chromosome Preparation 1. Blood culture. Heparinized peripheral human blood (1 ml) was added to 17 ml of prewarmed Iscove’s modified DMEM medium with 1% GPS (L-glutamine, penicillin, and streptomycin antibiotic mix). Two milliliters of fetal calf serum and 200 µl phytohemaglutinin were also added. The solutions were gently mixed in a flask, which was placed lying flat in an incubator for 72 h at 37°C. 2. Harvesting for metaphase chromosomes (chromosome suspension preparation). One hundred microliters of colchicine was added to the culture, mixed gently, and incubated for 1 h at 37°C. The culture was transferred to a centrifuge at 1 000 rpm for 10 min. The supernatant was removed and the pellet was resuspended in leftover fluid. Five milliliters of prewarmed 0.075 M hypotonic KCl solution was gently added by pouring it down along the side wall of test tube in a dropwise manner. The pellet was thoroughly resuspended in KCl and incubated for 20 min at 37°C. The sample was centrifuged again at 1 000 rpm for 10 min. The supernatant was discarded and the pellet was resuspended in 5 ml of freshly made fixative, 3:1 methanol:acetic acid. It was centrifuged at 1 000 rpm for 10 min, and the supernatant was discarded and resuspended in fixative again. This step was repeated two to three times until the pellet and supernatant became clear. At this stage, the chromosome pellet in the fixative solution was stored at 4°C or metaphase slides were prepared as below. 3. Slide preparation. Prior to the procedure, new slides were soaked in absolute ethanol and left in a 4°C refrigerator for a few minutes. A previously prepared fixed (in fixative solution, methanol:acetic acid at 1:3) cell suspension, stored at ⫺20°C, was centrifuged at 1 500 rpm for 7 min. The supernatant was discarded and the cell pellet was resuspended in leftover fixative solution to achieve the right concentration. Slides in absolute ethanol were removed individually and dried. A drop of suspension was dropped onto the slide. The slide was air-dried. Slides were checked under a phase-contrast microscope to ensure that cell density was correct and that there were enough well-spread, cytoplasm-free mitoses. 4. RNase treatment. RNase A (Boehringer) stock solution (20 mg/ml) was diluted 1:200 in 2⫻ SSC (0.15 M NaCl, 0.015 M sodium citrate), applied to the slide (coverslip was 24 ⫻ 60 mm2 ), and incubated for 40 min at 37°C. The slide was then washed in 2⫻ SSC for 5 min three times, with shaking at room temperature. 5. Protein digestion. Ten microliters of pepsin (100 mg/ml) was added to 100 ml of 0.01 M HCl (1 ml HCl in 99 ml dH2O). Slides were incubated in pepsin solution for 5 min at 37°C. The slides were washed in PBS buffer for 10 min at room temperature followed by a single wash in PBS/MgCl2 for 5 min. Finally, the
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slides were dehydrated in an alcohol series prior to analysis under a scanning force microscope. B. Scanning Force Microscopy A commercial scanning force microscope was used (D3100, Digital Instruments, Santa Barbara, CA, www.di.com). This scanning force microscope is equipped with an optical microscope and a motorized stage with micrometer precision to move the sample with respect to the tip. Both allow the precise location of the chromosome spreads on the slide. The measurements were taken in air with a relative humidity of 40–50% during the experiments. The scanning force microscope was operated in contact mode. No significance improvements were observed in the spatial resolution by using dynamic modes such as intermittent contact mode or noncontact mode. It should be noted that the chromosomes in air are dehydrated and are hard enough to be imaged in contact mode without any damage (Fritzsche and Henderson, 1997). V-shaped silicon nitride cantilevers were used with the lowest accessible normal spring constant, 0.06 N/m (nominal value). The tips were oxide-sharpened. The normal forces between the tip and the sample were kept below 20 nN. During the topography data acquisition, the frictional force between the tip and the sample was also measured. Differences in the frictional force were associated with compositional variations of the surface (Overney et al., 1992; Tamayo et al., 1996). III. RESULTS
A. Effect of the Treatment on the Substrate To evaluate the effect of the enzymes on the chromosome structure, topographic images of the regions between chromosomes (substrate) were taken. The flatness of the substrate allows the determination of the effect of both enzymes, RNase and pepsin, in the cell material adsorbed on the glass support. Figures 1a and 1b show the images of the glass support after chromosome deposition in two different preparations. Though the cell material adsorption on glass is different in both preparations, there are common features. The surface has a significant roughness of 2.5–2.7 nm in comparison to the glass (⬃0.3 nm). This indicates that cell material has been adsorbed on the glass support after chromosome deposition. The rough surface consists of diffuse protuberances with a characteristic size of about 200 nm. On the other hand, holes can be seen on the substrate surface. The holes are larger in Fig. 1a with a diameter of 100–150 nm and a depth of about 25 nm. The hole diameter and depth in Fig. 1b are about 50 and 20 nm, respectively. The effect of the RNA removal from the substrate is shown in Fig. 1c. The substrate showed a granular structure with a grain size of about 50 nm with the same holes as before RNase treatment. The change in the surface is a result of the removal of the diffuse protuberances of the untreated substrate. This suggests that these protuberances can be associated with RNA molecules. In fact, when protein digestion by pepsin is applied to the untreated substrate, the diffuse protuberances appear as well-defined is-
lands, indicating that these islands are composed mainly of RNA (Fig. 1d). Figure 1e shows the effect of protein digestion on the RNase-treated substrate shown in Fig. 1c. A glass support free of cell material is achieved. The glass identification is based on its well-known slight roughness. Some residual particles from the treatments appear adsorbed on the glass. The holes seen before protein digestion (Fig. 1c) disappear. If the holes in the substrate after RNA digestion go through the protein layer to the glass, the depth of these holes would be the protein film thickness. To examine this possibility, friction force experiments were performed in larger holes in the substrate after RNase treatment (data not shown). The frictional force between the tip and the sample was different on the bottom of some holes and the substrate, indicating different compositions (Overney et al., 1992; Tamayo et al., 1996). This suggests that the bottom of these holes is the bare glass and indicates that the protein layer thickness in this preparation is 29–32 nm. When protein digestion by pepsin is applied after chromosome deposition, two effects are visible on the substrate (Fig. 1d). First, the holes disappear. This is consistent with the removal of a protein layer covering the sample as was described before. The other effect is that islands in the same position as the diffuse protuberances in the untreated substrate cover partially the glass substrate. The islands are clusters of globular-like units with a diameter of 50–60 nm. The island height is 12–20 nm. Figure 1f shows the effect of RNase treatment on the pepsin-treated surface shown in Fig. 1d. The islands that covered the glass before RNA digestion surprisingly remain. However, the size of the islands decreased. The diameter of island subunits and the island height were reduced to 40–50 and 7–12 nm, respectively. Although protein digestion and RNA digestion were applied in both preparations (Figs. 1a and 1b), the order of the treatments has an influence on the removal of the cell material adsorbed after chromosome deposition. RNA removal followed by protein digestion removes most of the cell material adsorbed on the glass support (Fig. 1e). However, cell material removal remains incomplete when RNase treatment is applied after pepsin treatment. The origin of this effect will be discussed under Section IV. B. Effect of the Treatment on the Chromosome Structure Up to now, the effect of the RNA and protein digestions in the regions between chromosomes in standard cytogenetic preparations has been analyzed. The adsorbed cell material on the glass is
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FIG. 1. (a and b) A region between chromosomes for two different standard cytogenetic preparations. (c and e) The region in a after RNase treatment and following pepsin treatment, respectively. (d and f) The region in b after pepsin treatment and following RNase treatment, respectively. The Z range is 50 nm. Scan size is 3.6 ⫻ 3.6 µm2.
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removed after, first, RNase treatment and, second, pepsin treatment. The consequences of this treatment on the chromosome morphology are illustrated in Fig. 2, which shows the image of an untreated chromosome (Figs. 1a and 1b), after the RNase treatment (Figs. 1c and 1d), and after the subsequent pepsin treatment (Figs. 1e and 1f). The lefthand images show the overall chromosome structure and the right-hand images show higher resolution images of the chromosome surface. Figure 2a shows the characteristic chromosome topography in standard cytogenetic preparations. The average chromosome width and height are 1.4–1.5 µm and 98 nm, respectively. These values are in agreement with other SFM studies (Fritzsche et al., 1997). The higher resolution image shows a quite smooth surface without evidence of chromatin fibers (Fig. 2b). The effect of RNA removal is shown in Figs. 2c and 2d. The average chromosome height decreased to 70 nm. A change from a smooth to a rough surface is produced (Fig. 2d). Both results indicate that a significant amount of RNA material was covering the chromosome. This can be seen in the changes in the substrate surface after RNA digestion (compare Figs. 1a and 1c). The chromosome surface after RNase and subsequent pepsin treatments is shown in Figs. 2e and 2f. The chromosome shows a well-defined centromere and chromatids (Fig. 2e). The chromatids are not completely separated. In some regions the chromatids are bridged and in others appear almost separated. The chromosome width is unchanged after the treatment, 1.5 µm. This value is the limiting width that separates prophase chromosomes that have not split into chromatids from metaphase chromosomes with separated chromatids based on electron microscopy studies (Sumner, 1991). Therefore, this chromosome is in an intermediate state between late prophase and early metaphase in which the chromatids are separating. Variations in the average chromosome thickness of about 5–15 nm along the chromatids are also seen (Fig. 3a). Musio et al. (1994) observed that standard chromosomes showed a height pattern similar to the G-banding pattern; i.e., thicker chromosome regions correspond to G bands. The chromosome length and the centromere position suggest that the images shown in Fig. 2 correspond to chromosome 8. Indeed, some optical G bands of chromosome 8 can be identified with the thicker chromosome regions. However, several G and R bands are difficult to distinguish using the height criteria. The most evident R bands are located at telomeres. The telomere regions are clearly thinner than the rest of chromosome, about 15 nm lower than the identified G bands. Furthermore, the images suggest a telomere region
structure different from the rest of the chromosome. This structure is defined by a dense nucleus surrounded by looser chromatin (Fig. 3b). At high resolution, the chromosome surface shows a granular structure with a grain size of 45–55 nm (Figs. 2f and 3b). This size is in agreement with the diameter of the chromatin fiber, 30 nm. Furthermore, the chromosome surface is clear of cellular material that could lead to mistaken conclusions (see Fig. 1e). It is therefore reasonable to relate each grain to the chromatin fiber at the surface folding back into the chromatid body (Harrison et al., 1983; Adolph et al., 1986). The tip-sample convolution explains the larger size of the grains compared to the chromatin fiber diameter. In SFM, the horizontal dimensions of objects whose size is comparable to the tip’s radius (5–50 nm) are overestimated (Keller and Franke, 1993). The changes in the chromosome structure when the protein digestion is applied before RNase treatment are shown in Fig. 4. The chromosomes prepared by the standard cytogenetic procedure show a width and height of 1.4–1.5 µm and 115–120 nm, respectively (Fig. 4a). No evidence of chromatin fibers was observed. After protein digestion by pepsin, a rougher chromosome surface appears (Fig. 4b). The chromosome width and height remained approximately the same. Figure 4a shows the chromosome morphology after pepsin treatment followed by RNase treatment. The chromosome width increased to 1.7 µm and the height decreased to approximately 105 nm. In this case, no splitting of the chromosome into chromatids is seen. This is consistent with a late prophase stage of the chromosome. At high resolution, a well-defined granular structure was not observed as in the chromosome shown in Figs. 2e and 2f. In fact, the cell material digestion was incomplete when the RNase treatment was performed after the pepsin treatment. Substrate images showed nanometer-size islands partially covering the glass support (compare Figs. 1b, 1d, and 1f). IV. DISCUSSION
A. RNA and Protein Digestion The analysis of the changes in the substrate after each enzymatic treatment reveals that a protein layer with embedded RNA molecules covers the chromosomes in standard cytogenetic preparations. The RNA molecules appear as diffuse protuberances on the untreated substrate (Figs. 1a and 1b). The diffuse aspect indicates that the RNA is embedded in the protein film. There are two reasons to associate these protuberances with RNA molecules: (i) After RNA digestion, the diffuse protuberances disap-
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FIG. 2. Topographic images of the same metaphase chromosome after each treatment step at different magnifications. (a and b) The untreated chromosome. (c and d) The RNase-treated chromosome. (e and f) The RNase- and subsequent pepsin-treated chromosome. (a, c, and e) Scan size, 10 ⫻ 5 µm2. (b, d, and f) Scan size, 4 ⫻ 4 µm2. The force was kept below 20 nN. The Z range is 150 nm. The height offset is set to highlight the surface features hiding the lower regions of the chromosome surface. Therefore the chromosome height cannot be deduced from the images. The values are specified in the text.
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FIG. 3. Images of the chromosome shown in Figs. 2e (a) and 2f (b). This chromosome has been treated with RNase and subsequently with pepsin. In both images, the grayscale was inverted and the contrast was increased to reflect the changes in the chromosome thickness. The size and the centromere position indicate that it is chromosome 8. Scale bar is 1 µm.
peared (Fig. 1c). (ii) After protein digestion, the diffuse islands appear well defined and adsorbed on the glass (Fig. 1d). However, the islands covering the glass after pepsin treatment were not removed after RNA digestion, although their size was reduced (Fig.
1f). This suggests that some proteins are somehow protected by RNA during the protein digestion. Tentatively, these proteins could belong to polysomes (Christensen, 1994). In fact, the dimensions of the globular units that compose the islands after protein
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digestion are in agreement with the ribosome size determined by SFM (Fritzsche and Henderson, 1998). A change from a smooth chromosome surface to a rough surface is produced after RNA removal. This indicates that a significant amount of RNA covers the chromosomes. In fact, a chromosome height reduction of about 20 nm is produced after RNA removal (Figs. 2a and 2c). A decrease in chromosome height of 10–30 nm has been found in other experiments. These results suggest that clusters of RNA molecules with a thickness greater than 20 nm cover the chromosomes in standard cytogenetic preparations. This is consistent with the tendency of the ribosomes to aggregate to the chromosome surface in solution (Marsden and Laemmli, 1979). Holes can often be seen on the substrate in standard cytogenetic preparations. These holes remain after RNA digestion. Some of these holes are connected to the glass support, allowing the estimation of the thickness of the protein film in standard cytogenetic preparations. Thus, the untreated chromosome shown in Fig. 2a is covered by a protein layer with a thickness of approximately 30 nm. Measurements in holes of other samples indicate that the standard cytogenetic procedure involves chromosomes embedded in a protein layer with a thickness of 15–35 nm. B. Chromosome Structure
FIG. 4. Topographic images of a chromosome without treatment (a), after pepsin treatment (b), and after subsequent RNase treatment (c). Scan size, 7 ⫻ 7 µm2.
In standard cytogenetic preparations, the chromosome surface did not show evidence of chromatin fibers. However, after RNA and subsequent protein digestion a granular structure is observed on the chromosome surface (Figs. 2f and 3b). We have two reasons to associate each grain with the chromatin fiber folding back into the chromatid. First, the size of the grains is about 50 nm, in agreement with the chromatin fiber size. Second, the chromosome has been cleaned of cell residues, mainly RNA and proteins. Indeed, the experiments discussed in the last section indicate that a protein and RNA film with a thickness greater than 50 nm covers the untreated chromosome shown in Fig. 2a. Thus, the chromatin fibers were hidden by the cell material film, resulting in a relatively smooth chromosome surface. Other SFM studies have claimed to image grains in the chromosome surface with a size similar to that of the chromatin fiber (De Grooth and Putman, 1992). However, this granular surface has not been reproducible. Furthermore, these features were always suspected to be artifacts due to salt and cellular debris (Fritzsche et al., 1997, De Grooth and Putman, 1992). In fact, we have found some untreated chromosomes with a granular structure with a grain size of 50–100 nm. The grains could be tentatively associated with the chromatin fiber cov-
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ered by a very thin protein and RNA layer. However, the substrate showed a similar grain structure. Indeed, the protein film that covers the glass regions between chromosomes showed a granular structure with a grain size of about 50 nm (Fig. 1c). Furthermore, in scanning electron microscopy studies, a brief trypsin treatment was required to image individual chromatin fibers in standard cytogenetic preparations (Harrison et al., 1983, Sumner, 1991). A very rich chromosome structure is achieved after RNA and protein digestion. It has been stated that prophase chromosomes are longitudinally split in two chromatids. However, in 1991, Sumner showed images of late prophase chromosome without a split into chromatids. Here, high-resolution images of a chromosome in an intermediate stage between singlechromatid prophase chromosomes and a doublechromatid metaphase stage have been shown. A pattern of transverse chromatin bands bridging both chromatids can be seen in the topographic images. The study of the chromatid splitting process is significant in the elucidation of the condensation process of metaphase chromosomes. Further work will be undertaken to study the chromosome structure in different prophase and metaphase stages. A chromosome height variation of 5–10 nm along its long axis is observed (Fig. 3a). Several thicker regions correspond to G and Q bands, consistent with previous SFM observations (Musio et al., 1994). However, the correspondence between optical bands and chromosome structure does not always relate to the chromosome thickness criteria. G bands seem to fulfill the criteria of thicker chromosome regions and joined chromatids. Further work is in progress to relate the optical banding and the chromosome structure. The most evident R bands are at telomeres. These bands are thinner than the rest of the chromosome. Indeed, the long-arm telomere region is 17 nm thinner than the adjacent G band. However, the chromosome thickness variation in the internal positions is less than 10 nm. Also, a looser chromatin conformation can be inferred in the telomeres. This is consistent with the higher transcriptional activity of the R bands (Holmquist, 1992). C. Structure Nativeness A protein film with embedded RNA molecules covers chromosomes prepared by the standard cytogenetic preparation. The RNA digestion followed by protein digestion by pepsin removes most of the cell material covering the chromosome. However, this treatment can alter the chromosome composition and structure. The chromosome consists of DNA as well as proteins. The chromosomal proteins can be divided into histones (H1, H2A, H2B, H3, and H4) and nonhistone proteins. Histones are responsible for the assembly of DNA into a 30-nm chromatin
fiber. Nonhistone proteins participate in the higherorder organization of the chromatin fiber and are responsible for the overall chromosome morphology (Adolph et al., 1977; Paulson and Laemmli, 1977). Untreated chromosomes lose mainly part of histone H1 after ethanol/acetic acid fixation (Burkholder and Duczek, 1982). However, this does not seem to produce any chromatin fiber disruption (Harrison et al., 1983). The protein digestion by pepsin is the most critical step. Although protein digestion by pepsin is considered a mild treatment, we have not found biochemical data on its activity. The pepsin treatment produces the digestion of a protein layer covering the sample with a thickness of 15–35 nm. It is also reasonable to consider that some chromosomal proteins can be removed or altered. However, the chromosome showed a granular structure identified with chromatin fiber loops at the surface after RNase treatment and subsequent pepsin treatment. This structure is similar to the chromosome structure without protein extraction studied by transmission electron microscopy (TEM) (Adolph et al., 1986). Furthermore, TEM studies of histone-depleted chromosomes in solution showed a scaffolding structure surrounded by DNA loops with a diameter of 10–30 µm (Paulson and Laemmli, 1977). Clearly, these loops are not observed in this work. These observations indicate that histones in chromatin fibers are hardly affected by the treatment. Nonhistone proteins are more resistant to protease treatments than the histones (Adolph et al., 1977). The chromosomes after RNA and subsequent protein digestions showed a compact morphology. Imaging in solution also showed a compact structure (Tamayo and Miles, in press). This indicates that the main nonhistone proteins remain after the treatment. In more detail, the beginning of nonhistone removal produces a filamentous morphology to the chromatin due to separation and unfolding of the loops of the chromatin (Harrison et al., 1983). This kind of structure was not observed here, indicating that the nonhistone proteins responsible for the chromosome morphology remain after the treatment. Similar chromosome structures were obtained with a threefold pepsin concentration and double the incubation time. However, the used pepsin concentration and an incubation time slightly longer than 3 min are enough to remove the protein film. The RNA removal does not affect the chromosome structure (Adolph et al., 1977). However, the SSC solution in which the RNase is diluted extracts some nonhistone proteins (Burkholder and Duczek, 1982). Indeed, when the RNase treatment is applied after protein digestion, the chromosome width increases slightly, from 1.5 to 1.7 µm. This suggests some chromatin loosening due to the extraction of some
CLEANING OF CHROMOSOME PREPARATIONS FOR SFM
nonhistone proteins. However, an increase in chromosome width was not observed when the RNase was applied to the untreated chromosome, suggesting that the protein layer covering the chromosomes screens the effect of SSC. Then, the RNA removal step does not affect the chromosomal proteins when it is applied before protein digestion. V. CONCLUSIONS
The SFM provides topographic images with high spatial resolution. Horizontal and vertical resolutions of 1 and 0.01 nm are currently obtained, respectively. However, no new information about the chromosome structure has been obtained by SFM. Furthermore, most of the known information about the chromatin structure comes from optical microscopy. The optical spatial resolution is two orders of magnitude lower than that with SFM. However, it is sensitive to the chromosome subsurface avoiding the surface contaminants. In our opinion, removal of cell material is a prerequisite to resolve the chromatin structure by SFM. RNA removal and subsequent protein digestion by pepsin treatment provided the following results: (I) Standard cytogenetic preparations consist of chromosomes covered by a protein layer with a thickness of 15–35 nm. RNA molecules are embedded in this protein layer and adhere preferentially to the chromosome surface. (II) The glass support regions between chromosomes were free of cell material after treatment. A well-defined chromosome structure was obtained and the chromatin fiber loops were resolved. Individual chromatin fibers could not be resolved in the untreated chromosomes. (III) The chromosome structure between late prophase and early metaphase is shown. The chromosome was not split into chromatids. A pattern of bridges joining the chromatids was observed. (IV) The telomeres showed a differential structure. They were clearly thinner than the rest of the chromosome and showed a looser chromatin conformation. This work was supported by the European Union TMR program. REFERENCES Adolph, K. W., Cheng, S. M., and Laemmli, U. K. (1977) Role of nonhistone proteins in metaphase chromosome structure, Cell 12, 805–816. Adolph, K. W., Kreisman, L. R., and Kuehn, R. L. (1986) Assembly of chromatin fibers into metaphase chromosomes analyzed by transmission electron microscopy and scanning electron microscopy, Biophys. J. 49, 221–231. Binnig, G., Quate, C. F., and Gerber, Ch. (1986) Atomic force microscopy, Phys. Rev. Lett. 56, 930–933.
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