Label-Free Determination of the Cell Cycle Phase in Human

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Label-free determination of the cell cycle phase in human embryonic stem cells by Raman microspectroscopy Stanislav O. Konorov, H. Georg Schulze, James M. Piret, Michael W. Blades, and Robin F. B. Turner Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/ac400310b • Publication Date (Web): 22 Aug 2013 Downloaded from http://pubs.acs.org on August 26, 2013

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Analytical Chemistry

Label-free determination of the cell cycle phase in human embryonic stem cells by Raman microspectroscopy Stanislav O. Konorov,1,2§ H. Georg Schulze,1§ James M. Piret,1,3 Michael W. Blades2* and Robin F. B. Turner1,2,4*

1. Michael Smith Laboratories, The University of British Columbia, 2185 East Mall, Vancouver, BC, Canada, V6T 1Z4 2. Department of Chemistry, The University of British Columbia, 2036 Main Mall, Vancouver, BC, Canada, V6T 1Z1 3. Department of Chemical and Biological Engineering, The University of British Columbia, 2360 East Mall, Vancouver, BC, Canada, V6T 1Z3 4. Department of Electrical and Computer Engineering, The University of British Columbia, 2332 Main Mall, Vancouver, BC, Canada, V6T 1Z4

§ Equal contributions

*

Corresponding Authors:

Turner, Email: [email protected], Fax: 604-822-2114 Blades, Email: [email protected], Fax: 604-822-2847

ACS Paragon Plus Environment


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2 Abstract The cell cycle is a series of integrated and coordinated physiological events that results in cell growth and replication. Besides observing the event of cell division it is not feasible to determine the cell cycle phase without fatal and/or perturbing invasive procedures such as cell staining, fixing, and/or dissociation. Raman microspectroscopy (RMS) is a chemical imaging technique that exploits molecular vibrations as a contrast mechanism; it can be applied to single living cells non-invasively to allow unperturbed analysis over time. We used RMS to determine the cell cycle phase based on integrating the composite 783 cm-1 nucleic acid band intensities across individual cell nuclei. After correcting for RNA contributions using the RNA 811 cm-1 band, the measured intensities essentially reflected DNA content. When quantifying Raman images from single cells in a population of methanol-fixed human embryonic stem cells, the histogram of corrected 783 cm-1 band intensities exhibited a profile analogous to that obtained using flowcytometry with nuclear stains. The two population peaks in the histogram occur at Raman intensities corresponding to a 1-fold and 2-fold diploid DNA complement per cell, consistent with a distribution of cells with a population peak due to cells at the end of G1 phase (1-fold) and a peak due to cells entering M phase (2-fold). When treated with EdU to label the replicating DNA and block cell division, cells with higher EdU-related fluorescence generally had higher integrated Raman intensities. This provides proof-ofprinciple of an analytical method for label-free RMS determination in situ of cell cycle phase in adherent monolayers or even single adherent cells. Key words: cell cycle, human embryonic stem cells, DNA, non-invasive, Raman microspectroscopy, label-free analysis, chemical composition mapping, single cells, in situ


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3 Introduction The highly ordered process of replication and distribution of genetic material from one cell generation to the next is a key component of the cell cycle. The cell cycle is divided into two discrete phases: mitosis (M phase) as a cell separates all of its material and divides into two daughter cells. This is followed by a gap phase (G1) wherein cells acquire and produce the necessary components for the DNA replication phase (S). S phase is followed by a second gap phase (G2) during which further growth occurs and the cell readies for mitosis. M phase is of short duration (~1 h), G1 is highly variable and lasts from ~3 h in embryonic stem cells to several times that in somatic cells, S phase is of consistent ~8 h duration while G2 phase is again relatively short (~4 h). The cell cycle in human cells therefore lasts at least ~16 h and can be much longer depending on the cell type.1 For effective replication and distribution of genetic material, the events in each phase should be completed before proceeding to the next phase. This is accomplished via checkpoints for monitoring the integrity of DNA at the G1/S and G2/M borders.2 The cells can also respond to external stimuli, often choreographed through a cascade of intracellular phosphorylations2, 3 that allow coordination between cell cycle progression and differentiation.4 Given the fundamental importance and the complexity of the cell cycle, it is of interest to study or manipulate its molecular, structural, and regulatory events and these often require determination of the phase of the cycle. The latter is accomplished by labeling/staining cells to reveal the phase of the cycle. Physical fractionation or chemical arrest of phases can be used to produce synchronized cell populations. Amongst cell cycle synchronization methods are serum starvation, temperature reduction, mitotic shake-off, flow cytometry-based cell sorting or the administration of chemical agents such as colchicine to block mitosis or hydroxyurea to inhibit DNA synthesis.5 Staining often requires fixing before applying DNA-binding dyes such as Hoechst 33342 (bisbenzimidazole) or DAPI (4',6-diamidino-2-phenylindole) but live cells can also be labeled.5, 6 These methods used to identify the phase of the cell cycle result either in cell perturbation or death. For example, although many methods have been developed to



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4 synchronize cell populations, they generally suffer from not being sufficiently effective, unacceptably perturbing the cell’s physiology, and/or being toxic and hence not suitable for living cells.5 A non-invasive method capable of determining the phase of the cell cycle in situ would therefore be of considerable interest. Raman spectroscopy is based on inelastic light scattering that carries information about molecular vibrational modes and is therefore sensitive to structure, composition and environment; moreover, it can be used effectively in aqueous environments as water is a weak Raman scatterer and a weak absorber at the excitation wavelengths typically used.7,

8

It does not require exogenous chemical labels and is readily coupled with an

optical microscopy platform to yield imaging or mapping capabilities commonly referred to as Raman microspectroscopy (RMS), which has been used successfully for the chemical analysis, quantification, and imaging of cells,7, 9-13 as well as revealing aspects of the cell cycle.8, 13-18 However, these previous works have not demonstrated that the phase of the cell cycle can be determined for several single cells in situ, label-free, without synchronization, and based on cellular DNA content only. Neither have thresholds been investigated to demarcate the boundaries between cell cycle phases. Specifically, Swain et al.8 synchronized the cells in culture and measurements from several single cells were averaged in that work, as also done by Matthews, et al.13 and Pliss et al.14 Based on Raman-fluorescence hybrid microspectroscopy of cell nuclei, Pully et al.15 could distinguish between interphase and mitosis (which can be done by optical microscopy as well), but not between interphase subdivisions. Short et al.16 obtained spectra from suspended cells and nuclei extracted from cells. The work by Kang et al.17 exploits functionalized gold nanoparticles for organelle-targeted in vivo labeling. Finally, the work by Huang et al.18 pertains to yeast, but not mammalian cells and offers no quantitative information relating DNA content to cell cycle phases. Nevertheless, the work by all of these authors provides valuable insight into approaches that could be used to refine cell cycle measurements and to more accurately establish transition points between the phases. In general, Raman spectroscopy suffers from weak signal intensities except when specialized variants of the technique are employed.19-21 To compensate for the weak scattering, high excitation light intensities and/or long spectral acquisition times are used.


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5 Fortunately, it has been observed that neither of these procedures, within limits, compromises the viability of exposed living cells.22 Consequently, Raman spectroscopy has become a powerful tool for investigating several distinct macromolecular components simultaneously in fixed or living cells, including cells in situ. We present here proof-ofprinciple of a technique to determine the phase of the cell cycle based on RMS which is reagentless and non-perturbing to live cells, and retains information about the spatial distribution of cells in various parts of the cycle. Materials and methods Approach. Conventionally, the determination of the cell cycle is based on applying a fluorescent nuclear stain to the cells and analyzing using into a flow cytometer. Consequently, a number density distribution as a function of fluorescence intensity can be created that reveals the number of cells in different phases of the cycle. This distribution has as lower bound the fluorescence intensity corresponding to the nuclear material of a single cell in G1 phase and normally double that amount as its upper bound, corresponding to those cells in G2/M phase. With the use of appropriate calibration, Raman spectroscopy is quantitative,19 hence the amount of DNA in a nucleus can be determined by utilizing a suitable Raman band (or combination of bands) to achieve adequate selectivity. For example, the 783 cm-1 band is a composite nucleic acid band consisting of overlapping RNA and DNA pyrimidine ring vibrations and an overlapping DNA O-P-O symmetric stretch (see Table 1). In the absence of collagen interference, the 811 cm-1 band is indicative of RNA such that it can be used to calculate from the intensity of the 783 cm-1 band the DNA content only. Consequently, the integration of the composite 783 cm-1 nucleic acid Raman band intensities across the cell nucleus, along with a correction to remove contributions from RNA, should give a measure of the DNA content of the nucleus and thus provide an indicator of the cell cycle. This information can then be used to create a number density distribution

as

a

function

of

Raman

band

intensity,

analogous

to

the

fluorescence/cytometry approach, or to assess the phase of the cell cycle in situ for individual cells. Indeed, in previous work, we found that the 783 cm-1 nucleic acid Raman band diminished in intensity by approximately half upon non-specific



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6 differentiation of hESCs.22 This finding was consistent with the much-shortened G1 phase and comparatively longer S phase in hESCs compared to differentiated progeny.1 Thus the higher nucleic acid content deriving from more cells being in S phase was reflected in the 783 cm-1 band. Cell culture. CA1s cell line was adapted for enzymatic dissociation23 from the CA1 line24 of hESCs generously provided by Dr. Andras Nagy (Mount Sinai Hospital Toronto, ON Canada). The cells were maintained in Matrigel-coated (BD Biosciences, Mississauga, Canada) culture dishes (Sarstedt, Montreal, Canada) with mTeSR1 medium (STEMCELL Technologies, Vancouver, Canada) to maintain pluripotency.

For

maintenance cultures, the medium was changed daily and the cells were passaged when the dishes approached ~80% confluency. For cultures used in cell cycle measurements, the medium was replaced, according to the manufacturer’s protocol, for 5 h with medium containing 5x the stock concentration of a modified nucleoside, 5-ethynyl-2´deoxyuridine (EdU), that is incorporated into DNA during DNA synthesis, (Click-iT EdU Imaging Kits, Molecular Probes, USA) to identify cells in S phase. Thereafter cells were methanol fixed. Fluorescence Imaging. Differential interference contrast (DIC; also known as Nomarski) images and Raman spectra were obtained from methanol fixed samples. Subsequently, samples were treated with reagents in the Click-iT® EdU Alexa Fluor® 647 Imaging Kit to visualize cells in S-phase of the cell cycle (red fluorescence) and simultaneously counterstained with Hoechst 33342 (Molecular Probes, USA) to visualize cell nuclei (blue fluorescence). The use of fixed cells in this study was dictated by the need to corroborate the Raman results with conventional fluorescence images, it is not required for future implementations of the method. Table 1 Raman band assignments of interest for human embryonic stem cell spectra. An extensive selection of band assignments and a collection of cytologically relevant Raman spectra can be found elsewhere.25, 26 Raman Assignment band 718 cm-1 Lipids (phospholipids  C-N stretch); DNA/RNA (adenine  ring breathing) -1 724 cm DNA/RNA (adenine  ring breathing)


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7 746 cm-1 757 cm-1 783 cm-1 811 cm-1 827 cm-1 853 cm-1 874 cm-1 937 cm-1 1003 cm-1 1031 cm-1

Thymine (ring breathing) Proteins (symmetric ring breathing in tryptophan) 782 cm-1: DNA/RNA (pyrimidines  ring breathing); 788 cm-1: DNA (backbone  O-P-O stretching) RNA (backbone  O-P-O stretching); proteins (collagen  C-C stretching of proline and hydroxyproline) Proteins (proline, hydroxyproline, out-of-plane ring breathing in tyrosine); DNA/RNA (asymmetric O-P-O stretching) Proteins (collagen  C-C stretch in proline; also other proteins  ring breathing in tyrosine); carbohydrates (glycogen, polysaccharides  C-O-C stretching) Proteins (collagen  C-C stretching in hydroxyproline, proteins  ring deformation in tryptophan); lipids (phospholipids  asymmetric stretch of choline); carbohydrates (C-O-C stretching) Proteins (collagen type I  C-C stretching, α-helix  C-C stretching); carbohydrates (glycogen) Proteins (symmetric ring breathing in phenylalanine) Proteins (collagen, keratin, C-N stretching in proteins, C-H in-plane bending of phenylalanine); lipids (phospholipids); carbohydrates (polysaccharides)

Raman Imaging. For Raman spectroscopy, 12.5-mm diameter glass-encapsulated (100-nm thick) gold mirrors (ThorLabs Inc, Newton, USA) were placed in polystyrene Petri dishes and coated with a solution of Matrigel diluted 1/30 in DMEM/F12 (Invitrogen) immediately followed by a 1-h incubation at room temperature. Undifferentiated hESC were seeded onto the coated mirrors and allowed to grow to high density (5 days) before methanol fixation as described above. Spectra were obtained at room temperature from these colonies using a Raman microspectroscopy system in “StreamLine” mode (InVia, Renishaw, Gloucestershire, UK) using a 50x objective lens. The gold mirror increased the Raman signal almost 4 times compared to a standard glassbottom Petri dish because both forward and backward-scattered signals from a two-pass beam path were collected while the encapsulating glass layer was too thin to generate a significant background.20 Raman scattering, generated with 100 mW of 785-nm excitation at the sample, was collected for 20 s per spectrum (i.e., from each pixel). An area of 180 µm x 200 µm was scanned with a step size of 1.1 µm in the horizontal direction and 1.1 µm in the vertical direction (with reference to Figure 1). The spectral range selected for use with this method (687 cm-1 - 1074 cm-1) was based on our previous work with hESCs.27-31 This range includes enough of the fingerprint region to



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8 permit quantitative univariate determination of DNA content while reducing the required signal acquisition time. Though not critical for the demonstration presented here, it is important for applications of this approach involving live cells to minimize exposure time and the related stress on the cells. While other DNA bands outside of this range can, in principle, provide additional information, they also include contributions from other species with overlapping bands and we have found that the net result does not yield a significant improvement in precision. Data Processing. Matlab 7.0 (The MathWorks, Natick, MA) was used for all spectral processing and data analyses. Spectral processing consisted of baseline flattening with a moving average, peak stripping, automated procedure,32 automated cosmic spike removal, and automated smoothing.33 This was followed by principal component analyses (PCA). Based on the processed Raman spectra, chemical contrast images and principal component images were generated, and further analyses performed, using custom written software. Raman-based images were used directly (e.g. Figure 1(a)) for quantitative purposes but also two-dimensionally interpolated with a spline function (2D spline) to aid in visualization (e.g. Figure 1(b)). Safety considerations. Cells are considered Risk Group 2 biological material and were handled according to Containment Level 2 procedures. All chemicals were handled according to the relevant manufacturer’s protocols and material safety data sheet instructions. Raman spectroscopy employed a Class 3b laser and the appropriate radiation safety procedures were followed. Results and discussion Nuclear boundaries needed to be discerned accurately in order to obtain a satisfactory estimate of the DNA based on integration of the 783 cm-1 band intensity, particularly since chromatin tends to concentrate around the periphery. But, because cells remained in situ during the analysis, it was on occasion difficult to correctly demarcate nuclear boundaries. To ascertain that Raman images permitted accurate nuclear or cellular level segmentation, we investigated several approaches based on: individual bands, ratios of bands, spectral principal component (PC) scores, correlation coefficients between cellular spectra and the spectrum of a compound of interest (e.g. RNA), and higher resolution


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9 scans (which produce larger data sets and take more time to cover a given area). Of these, we generally found images based on the second PC (PC2) scores of Raman spectra (Figure 1(b), 2D interpolated) the easiest to work with as nuclei appeared well demarcated. A Raman image (not interpolated) of a scanned hESC colony area is shown in Figure 1(a) for the 811 cm-1 band. The inset is a reduced DIC image of the same area. Fluorescently stained nuclei of the same area (Figure 1(c)) are shown bright green to provide contrast with the red/orange nuclei in the overlaid PC2 score map (Figure 1(d)). Differences in optical paths to separate cameras used for Raman and fluorescence measurements caused small misalignments between nuclei in these overlaid images. Artifacts due to staining or necrotic nuclei were apparent as small, irregular bright spots (indicated by arrows). Nuclear boundary ambiguities in the PC2 images (e.g. black rectangle) were resolved by reference to fluorescence (e.g. red rectangle), DIC, and/or combination DIC/Raman images (example image in Figure 3(c)). Using the PC2-based image (Figure 1(b), but without interpolation), the 783 cm-1 Raman band intensities were integrated for all the spectra within a nucleus (e.g. those shown enclosed in black polygons in Figure 2(a)) and the same was done for the 811 cm-1 Raman band intensities. Because, for RNA, the 811 cm-1 band is approximately half of the 783 cm-1 band (Figure 2(a), RNA spectrum inset), the RNA contribution to the 783 cm-1 band intensity could be adjusted by subtracting twice the integrated 811 cm-1 band intensities from the integrated 783 cm-1 band intensities. From the 40 arbitrarily chosen cells (not all visible in Figure 2(a)), the histogram in Figure 2(b) was created. The histogram shows the number of cells with integrated 783 cm-1 Raman band intensities, corrected for RNA content, that fell within continuous discrete bins along a continuum of 783 cm-1 Raman band intensities. This constituted a discrete approximation to the quasicontinuous distributions familiar to users of flow cytometry methods.



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Figure 1 (a) Shown are 811 cm non-interpolated Raman image (high intensities in red, low intensities in blue) of a hESC colony area (inset: reduced DIC image of same area) and corresponding (b) spline-interpolated second principal component score-based Raman (high intensities in red, low intensities in blue) and (c) nuclear stained fluorescence images. (d) Overlay of Raman image in (b) on fluorescence image in (c) reveals generally matching nuclear boundaries; small differential optical path-dependent mismatches occurred due to separate fluorescence and Raman cameras. Ambiguities in Raman image nuclear boundaries (e.g. black rectangle) could be resolved by reference to fluorescence (e.g. red rectangle) and/or DIC and/or other Raman images (see text). Small, irregular, bright green dots of fluorescence image (e.g. black arrows) are necrotic cell nuclei/artifacts of staining. -1


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11

Integrated intensities for the 40 analyzed cells ranged from ~ 11,000 to ~ 43,000 arbitrary units. The first peak occurred around 21,000 and the last peak around 42,000 arbitrary units. We therefore considered the first peak (Figure 2(b), red bars) to represent cells in the G1 phase, while the last peak (Figure 2(b), green bars), with approximately double the intensity of the first one, represented cells in G2/M phase. We used a threshold of 23,500 arbitrary units to demarcate the upper boundary of the G1 phase and 36,000 arbitrary units for the lower boundary of G2/M phase. Cells between these limits (Figure 2(b), blue bars) were considered to be in S phase. Given the rapid cycling of nonconfluent hESCs,1 a large fraction (17/40) of S-phase cells was expected.

Figure 2 (a) Map based on the PC2 scores of Raman spectra collected from a scanned hESC colony. 40 cell nuclei (not all visible) in the area scanned were identified and encircled (black polygons) and the 783 cm-1 band intensities for the spectra corresponding to the map locations within each polygon were integrated. The inset shows the mean spectrum of the selected nuclei and the spectrum of RNA. The RNA peak at 811 cm-1, quite weak in the mean spectrum, was used to make a correction in order to obtain an improved estimate of the DNA-only content in the polygon-enclosed regions. (b) A 20-bin histogram of DNA-only content from the cells in (a) showing the number of cells within each bin. Based on their individual integrated peak intensities, cells were considered to be in G1 phase (red), S phase (blue), or G2/M phase (green). To assess whether we could indeed identify cells in S phase based on corrected integrated 783 cm-1 band intensities, Raman and fluorescence measurements were made



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12 on EdU-treated cells. During S phase, EdU is incorporated into nuclear DNA and can subsequently be visualized with specific fluorescent antibodies.34 In Figure 3(a), a DIC image is shown of an area of a colony treated with EdU to identify cells in S phase. Raman spectra obtained from scanning the same area were used to construct a 2D splineinterpolated image based on the 811 cm-1 RNA band (Figure 3(b)). Because RNA was primarily concentrated in the cytoplasm,35 nuclei were visible as low-intensity (blue) disks. This image was quite similar to 811 cm-1 RNA band images from untreated colonies (Figure 1(a)) suggesting that EdU did not materially alter the utility of Raman images. In DIC images nuclei appear as flat disks bordered by ridges and in 811 cm-1 Raman images as blue disks bordered by yellow/orange/red areas. An overlay (Figure 3(c)) of the DIC image on the 811 cm-1 Raman image revealed good correspondence between the DIC and Raman images and we used it to help resolve ambiguities about nuclear boundaries. An image of fluorescent EdU antibodies (Figure 3(d)) allowed us to identify those cells in this area that were in S phase during EdU treatment. The fluorescence image of Figure 3(d) was superimposed on the PC2 score-based image from the scanned area (analogous to Figure 2(a)) causing EdU-containing cells to appear yellow/brighter than non-EdU-containing cells (Figure 3(e)). The PC2 image was used to retrieve all the spectra from a given cell nucleus (i.e. spectra coming from within regions enclosed by black polygons) for a number of cells clearly in S phase (labeled 1 to 7; EdU+) and cells not in S phase (labeled 8 to 13; EdU-) or cells possibly entering S phase as the experiment ended. Artifacts precluded the use of all scanned cells. As expected, the RNA-corrected, integrated 783 cm-1 Raman band intensities (Figure 3(f)) revealed that EdU+ cells had generally higher DNA content than EdU- cells. When using the threshold of 23,500 arbitrary units to demarcate the upper boundary of the G1 phase (see transition between red/blue bars in Figure 2(b); blue line Figure 3(f)) a majority of EdU- cells fell into the G1 phase category while a majority of EdU+ cells fell into S phase category. A Kolmogorov-Smirnov two-sample directional test on integrated 783 cm-1 Raman band intensities revealed a statistically significant difference between the two groups when RNA-corrected (EdU- < EdU+; test statistic = 0.67; p < 0.05), but not in the absence of RNA correction. Thus, the Raman intensities of EdU- cells were consistent with having been in G1 phase when compared to the intensities of S phase EdU+ cells.


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Figure 3 hESCs were treated with EdU to identify cells in S phase. (a) DIC image of a colony area, (b) Raman image of the same area based on the 811 cm-1 RNA band was similar to those from untreated colonies suggesting that EdU treatment did not materially alter the utility of Raman images (more below). (c) DIC image superimposed on 811 cm-1 Raman band image shows that RNA predominantly accumulated in the cytoplasm leaving nuclei appear dark blue. This image was used to help disambiguate nuclear boundaries. Vertical stripes are artifacts. (d) Immunostained fluorescence image showing EdU incorporation into DNA, hence cells in S phase, of the same section. (e) Fluorescence image superimposed on PC2 scores map (analogous to Figure 2(a)) caused EdU-stained cells in S phase to appear yellow. The 783 cm-1 Raman band intensities were integrated (and RNA-corrected, see Figure 2) for seven moderate to heavily stained cells (1 - 7) and a further six weakly stained or unstained cells (8 - 13) to compare Raman estimates of the S phase to EdU results. (f) Weakly stained or unstained cells had lower integrated RNA-corrected 783 cm-1 Raman band intensities (disks) compared to moderately or heavily stained cells. Without RNA-correction (squares), the difference was not significant. Blue line indicates G1/S cutoff used in Figure 2(b). (g) Some evidence of spectral distortion was evident when comparing a colony of EdU-treated (EdU+) cells to a colony of cells not treated (EdU-). Nucleoli also appeared prominent in images after EdU treatment (arrows in panel (c)).



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We also found that EdU treatment caused visible structural and spectral changes in cells. Compared to non-treated cells, nucleoli appeared more pronounced (‘blister-like’) within nuclear disks (arrows, Figure 3(c); also compare Figure 3(a) to the inset of Figure 1(a)) while subtle changes emerged in the Raman spectra (Figure 3(g)). During S-phase, cells incorporate EdU into nuclear DNA. Thus, EdU treatment reveals which cells are in S-phase, but it does not synchronize cells in S-phase unless treatment is prolonged. One would therefore not expect to see substantial differences in spectra averaged over an EdU treated colony and a similar one not so treated because cells would, on average, have similar phase distributions in both. The subtle differences in peaks (g) therefore reflect the effects of EdU, not the difference between S-phase and other phases. As cells enter Sphase, they will have (corrected) 783 cm-1 Raman intensity values close to those in G1 phase. When cells exit S-phase, they will have intensity values close to those in G2 phase. These spectral differences are not subtle, as evident from Figure 2(b) where G1 phase cells have integrated Raman intensity values around 21,000 arbitrary units while G2 phase cells have approximately double that. We suspected that the diminished 757 cm-1 band could have been due to reduced contributions from the nearby 746 cm-1 thymidine ring breathing mode26 and the slight increase and downshift in the 783 cm-1 band could have been due to altered uracil ring breathing contributions.26 Although both interpretations were consistent with the substitution of EdU for thymidine, further analysis would be needed for confirmation. Nevertheless, these observations suggest that some caution is warranted if a quantitative interpretation of Raman and fluorescence data from EdU-treated cells is needed. Amongst other difficulties were occasional remaining ambiguities about nuclear boundaries in Raman images that tended to increase the uncertainties in the integrated 783 cm-1 band intensities. We found that we could improve the location of nuclear boundaries by increasing the spatial resolution of scans, but at the cost of increased measurement time. Also, the RNA 811 cm-1 band could be conflated with an overlapping band from collagen thus requiring a more complex correction to be devised. Although this possibility exists in principle, a significant collagen confound overlapping the nucleus is unlikely for most cell types. Finally, for large numbers of cells, this mode of


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15 analysis is time consuming owing to the spectral acquisition and processing requirements, but we expect continued technological advances to ameliorate this limitation. Although we have used fixed cells for the proof-of-principle data presented here, this technique can, in principle, be used to investigate live cells. Its application to live cells has in fact been our goal and part of the motivation for this proof-of-principle investigation. For some time we have been interested in the analysis of live cells in situ to understand niche formation and the emergence of inhomogeneities in stem cell populations as these affect their differentiation. Accounting for the effects of cell cycle variations has been problematic and the greatest source of uncertainty in such analyses However, applications involving live cells are subject to an inherent limitation of using near-infrared spontaneous Raman microscopy in that long image acquisition times are necessitated by the need to keep the laser power density at the sample below the damage threshold for live cells (ca. 100 mW). This severely limits the number of cells that can be analyzed since the total aquisition time must remain negligible compared to the cell cycle period. This limitation might be addressed by, for example, developing an approach based on locating cell nuclei using DIC or phase-contrast images and obtaining Raman spectra from the nuclei only. This reduced scan area, or scanning with lower resolution/larger spot size, would reduce scanning times enough to permit the analyses of more live cells in a colony. Importantly, the approach presented here could also be used with stimulated Raman scattering microscopy, which would reduce the acquisition time to a fraction of what is needed with spontaneous Raman microspectroscopy. Conclusions The cell cycle is a series of coordinated physiological events of fundamental importance for cell replication. During this process, the genetic material of a cell is doubled, increasing Raman scattering from DNA-related bands. By integrating the composite 783 cm-1 nucleic acid band across cell nuclei, and correcting the result for its RNA contributions using the RNA 811 cm-1 band, we confirmed that Raman intensities, analogous to fluorescence intensities in flow cytometry, could be used to determine the phase of the cell cycle. Importantly, we could do this for individual cells in situ, thus



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16 preserving potentially valuable contextual information, including local aspects of colony morphology such as niche-formation.27 We believe that our approach has tremendous potential for some applications. In particular, it offers researchers the option to study the cell cycle in its local environmental context, without the need to synchronize or label or otherwise perturb cells. At present, there is no other method that offers these particular advantages.

Acknowledgements We gratefully acknowledge Chris Sherwood for assistance with all cell culture related work. Funding was provided by the Natural Sciences and Engineering Research Council of Canada, the Canadian Institutes of Health Research, the Canada Foundation for Innovation, and the British Columbia Knowledge Development Fund.


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Label-Free Determination of the Cell Cycle Phase in Human

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