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Imaging Mass Spectrometry of Three-Dimensional Cell Culture Systems Haohang Li and Amanda B. Hummon* Department of Chemistry and Biochemistry, University of Notre Dame, 251 Nieuwland Science Hall, Notre Dame, Indiana 46556, United States
bS Supporting Information ABSTRACT: Three-dimensional (3D) cell cultures have increased complexity compared to simple monolayer and suspension cultures, recapitulating the cellular architecture and molecular gradients in tissue. As such, they are popular for in vitro models in biological research. Classical imaging methodologies, like immunohistochemistry, are commonly used to examine the distribution of specific species within the spheroids. However, there is a need for an unbiased discovery-based methodology that would allow examination of protein/peptide distributions in 3D culture systems, without a need for prior knowledge of the analytes. We have developed a matrix-assisted laser desorption/ionization-mass spectrometry (MALDI-MS)based imaging approach to examine protein distributions in 3D cell culture models. Using colon carcinoma cell lines, we detect changes in the spatial distribution of proteins across 3D culture structures. To identify the protein species present, we are combining results from the MS/MS capabilities of MALDI-MS to sequence peptides in a de novo fashion and nanoflow liquid chromatographytandem mass spectrometry (nLCMS/MS) of homogenized cultures. As a proof-of-principle, we have identified cytochrome C and Histone H4 as two of the predominant protein species in the 3D colon carcinoma cultures.
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hen cultured under appropriate conditions, many cell lines assume three-dimensional (3D) structures that mimic cellular architecture found in the body.1 These cell culture systems fill a valuable niche in experimental research; they blend the flexibility of cell culture systems with the ability to assume more complicated cellular structures and morphologies than are possible in simple two-dimensional (2D) monolayer or suspension systems. Some cellular properties, for example, polarity of epithelial cells, can be recapitulated in 3D systems but not in simpler 2D systems.2 In addition, it has previously been shown that gene expression patterns for these structures more closely resemble naturally occurring patterns in tissue than simple 2D cultures.3,4 3D culture systems are also more cost-effective and less time-consuming than using animal models. As such, 3D culture systems have been successfully used to address a variety of research applications: high-throughput drug screening,5 bioreactors for tissue engineering,6 studying hostpathogen interactions,7 and extravasation processes in metastases.8 Besides modeling the structural architecture, 3D culture models display some of the complex molecular gradients found in tissue. For example, the immortalized human breast cell line, MCF-10A, form acini structures, hollowed spheres of polarized cells that mimic mammary ducts.1 The acini take ∼20 days to grow in culture and show distinct patterns of protein expression and spatial distribution at each stage of growth. Many colon carcinoma cell lines form rounded heterogeneous structures, called spheroids, which mimic microtumors when grown to sufficient size (Figure 1a,b). When a spheroid exceeds ∼500 μm r 2011 American Chemical Society
in diameter, gradients for oxygen, nutrients, and catabolites develop within the structure.9 Cellular responses reflect these gradients. Robust, peripheral cells are highly proliferative and resemble actively cycling cells in vivo. The innermost cells become quiescent and form a necrotic core due to the deprivation of metabolites and oxygen.1,10 Differences in a variety of cellular responses are observed, including increased DNA strand breaks and increased lactate accumulation in the core. The outermost cells have higher glucose and ATP concentrations compared to the oxygen-deprived core.9 Imaging mass spectrometry (IMS) is an application of matrixassisted laser desorption/ionization mass spectrometry (MALDIMS), which utilizes soft-ionization techniques, enabling determination of the identity and location of molecules within a tissue sample.11,12 MALDI-IMS has proven to be a valuable technology with numerous publications describing its application in profiling proteins in diseased versus normal states,13,14 identifying lipid distributions on cell membranes,15,16 examining the locations of drugs and their metabolites,17 and mapping neuropeptides in the brain.18,19 MALDI-IMS has often been described as molecular histology, because of the wealth of unbiased information on molecular components that can be obtained.20 Secondary ion mass spectrometry (SIMS) and desorption electrospray ionization (DESI) mass spectrometry are complementary imaging approaches to MALDI-IMS. Compared to Received: September 5, 2011 Accepted: October 12, 2011 Published: October 12, 2011 8794
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Figure 1. (a) An optical image of a colon carcinoma spheroid after 10 days of growth. Prepared by seeding ∼6000 HCT 116 cells on an agarose meniscus in a 96-well plate. (b) Comparison of a spheroid (white dot near Lincoln’s nose) against a penny. The penny is placed on a centimeter ruler for scale. (c) A photograph of a spheroid at the tip of a 2 mL serological pipet. The arrow points to a spheroid. (d) An optical image of an ITO slide with four consecutive 10 μm thick slices of gelatin and spheroids prepared in a single well of a 24-well plate. The locations of four spheroids (with four consecutive slices apiece) are highlighted with arrows. Up to 16 different spheroids can be sectioned on each ITO slide or multiple consecutive slices of up to four spheroids.
MALDI or DESI imaging, SIMS has much higher spatial resolution, with pixel sizes in the single micrometer range.21 This spatial resolution enables subcellular imaging not possible with MALDI or DESI approaches22 and is readily adaptable to three-dimensional profiling.23 SIMS has been used to image lipids,24 proteins,25 and pharmaceuticals.26 One disadvantage is that SIMS cannot identify species via MS/MS. DESI-IMS has similar spatial resolution to MALDI-IMS. However, unlike MALDI-IMS, DESI is not limited by the interference of matrix ions in the low m/z range, making it an ideal approach for the analysis of lower molecular weight species.21 For example, DESI-IMS is well suited for the analysis of pharmaceuticals27 and explosives.28 Similar to MALDI and SIMS, DESI can be used to detect lipids29 in tissue sections, and it also allows sequential detection of proteins by MALDI on the same tissue section.30 While IMS provides information on the spatial location of analytes, it is equally important to determine their identity. Common methods to identify proteins in biological samples include nanoflow liquid chromatographytandem mass spectrometry31 (nLCMS/MS) and sequencing via MALDI.32 In typical
nLCMS/MS experiments, proteins are harvested from the sample of interest, fractionated either via liquid chromatography or one-dimensional gel electrophoresis, and then digested by trypsin or another endoprotease. The complex mixture of peptides is reduced and alkylated and again separated by nanoreverse phase liquid chromatography prior to analysis by electrospray tandem mass spectrometry.33 Peptide sequencing by MALDI can be accomplished via postsource decay (PSD)34 or in-source decay (ISD)35 with samples ranging from spotted mixtures of digested peptides to intact tissues. In PSD, fragments formed between the ion source and the reflector are used to determine the sequence.32 For ISD, fragmentation occurs in the source during a delay in the extraction time.32 Tissue slices are frequently analyzed by IMS. There are numerous reasons to examine tissue slices by IMS, including differentiating tissue subtypes36 and comparing the mass spectrometric results with those generated through classic histology approaches.37 However, there are limited perturbations that can be applied to tissues, especially when considering human material. In addition, the complexity of in vivo samples can make it difficult to control the molecular 8795
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Analytical Chemistry environment. For these reasons, cell culture systems are critical to address certain fundamental questions in biology, with 3D cultures playing an important role. While MALDIIMS has been extensively used to examine tissue slices, the application of the approach to 3D culture systems has not been reported. The ability to examine the spatial distributions of proteins within these cell complexes would expand the utility of 3D cell culture models in biological research. We have shown that the spatial distribution of analytes within these structures can be mapped via MALDI-IMS, with specific species detected only in the central region of the cultures and other species present throughout the structures. In addition to mapping the location of species, we have also demonstrated that individual proteins can be identified through a combination of MALDI sequencing and nLCMS/MS.
’ EXPERIMENTAL SECTION Cell Culture Plate Preparation. Spheroids were prepared in flat-bottomed 96-well microtiter plates (Thermo, Rochester, NY) according to the protocol described by Friedrich et al.10 Agarose (0.15 g) (Sigma-Aldrich, St. Louis, MO) was added to 10 mL of McCoy’s 5A cell culture media in an autoclave flask. The flask was sealed and autoclaved for 30 min at 120 °C, 200 kPa. The flask was placed in a water bath, at 60 °C, to prevent solidification of the agarose solution. The flask and the water bath were moved to a sterile benchtop. A volume of 50 μL of the agarose solution was added to the inner 60 wells of the 96-well microtiter plate with a 501000 μL automatic pipet. Multichannel pipettors were not used because they often produce air bubbles in the agarose meniscuses. The 36 outer wells in the plate were not used as they have higher media evaporation rates. The agarose solidified in ∼30 s after being transferred into the well. The plate was covered, and the agarose was allowed to cool down to room temperature. The covered plate was then stored in a refrigerator (∼4 °C). The plate should be used within 1 week of preparation. Cells. HCT 116 colon carcinoma cells (ATCC, Manassas VA) were used to grow spheroids. The tumor cells were subcultured for at least 4 but no more than 20 passages before the initiation of spheroids. The cell cultures were grown in a humidified environment with 5% CO2 at 37 °C. The cells were passed every 4 days. Cell lines were used within 3 months after receipt or resuscitation of frozen aliquots thawed from liquid nitrogen. The provider assured the authentication of these cell lines by cytogenetic analysis. Seeding the Cells. Cell suspensions were prepared by mild enzymatic dissociation using a 0.25% trypsin solution (Invitrogen, Carlsbad, CA). The concentration of the cell suspensions was evaluated via hemocytometer. The suspensions were diluted with cell culture media to ∼30 cells/μL. A volume of 200 μL of this suspension was transferred to each of the wells in the agarose-coated cell culture plate. Multichannel pipeters could be used to accelerate this seeding process. The number of cells in each well was approximately ∼6000. Incubation. The plates were incubated in a humidified environment with 5% CO2 at 37 °C to allow the spheroids to grow. The cell culture media was replenished every 35 days, depending on the growth rate. The old media were aspirated with a glass pipet connected to a vacuum pump; the pipet tip was kept at the interface of air and media so that the negative pressure did not remove the spheroids. After 2 weeks incubation, the spheroids
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were transferred to 24-well microtiter plates for further growth. The protocol to prepare agarose-coated 24-well plates was the same as for the 96-well plates. Gelatin Assisted Sectioning (GAS). When the spheroids grow to 1 mm in diameter (1420 days in culture for 6000 initial numbers of cells), they are ready for analysis. Media was aspirated and the spheroids were washed with phosphate buffered saline (PBS). The tumor spheroids were transferred to the wells of gelatin-coated 24well microtiter plate. The concentration of gelatin used was 350 mg/ mL. The gelatin solution should be prepared in a water bath, at ∼70 °C. The coating protocol is the same to prepare an agarosecoated plate. The gelatin solution tends to form a meniscus in the well of a 24-well plate because of surface tension, leaving a curvature after solidification. To minimize the curvature of the meniscus, a small amount of gelatin solution was dropped in the center of the meniscus. A serological pipet was used to transfer the spheroids. The media containing the spheroid was extracted via a seratological pipet, and the pipet was placed at an angle to the benchtop to allow the spheroid to subside to the tip of the pipet (Figure 1c), then the tip of the pipet was allowed to contact the gelatin coating (supporting layer) in a 24-well plate. The supporting layer can hold up to five spheroids without overlapping. The spheroids were covered with a thin later of warm gelatin (blanket layer). The blanket layer was allowed to solidify at room temperature. The solidified gelatin-embedded spheroids pellet can be scooped out with a spatula. The pellets were sectioned into ∼10 μm thick slices with a low-temperature microtome at 30 °C. The slices were thaw-mounted onto indium tin oxide (ITO) coated glass slides. One ITO slide can hold up to ∼34 slices or 1216 spheroids sections (Figure 1d). Slide Preparation. Optimal spectral quality is obtained if the sample is analyzed immediately after the sample preparation. Washing the slices with 70% isopropanol followed by 95% isopropanol extends the lifetime of the samples.38 The washing was carried out by gently submerging the slide in the isopropanol solutions for less than 10 s. Matrix Application. 2,5-Dihydroxybenzoic acid (DHB) (SigmaAldrich, St. Louis, MO) and sinapinic acid (Sigma-Aldrich, St. Louis, MO) were used as the MALDI matrixes. The DHB matrix was prepared by dissolving DHB in 50:50 methanol water with 0.1% trifluoroacetic acid (TFA) to yield a concentration of 30 g/L. The sinapinic acid matrix was prepared by dissolving sinapinic acid in 50:50 acetonitrilewater with 0.1% TFA to yield a concentration of 30 g/L. The DHB matrix was distributed onto the sample using a pipet. Sinapinic acid matrix was applied by airbursh or pipetting a 200 nL volume. The matrix was allowed to dry between intervals. Matrix application was monitored with an optical microscope with 10 magnification. The slides were dried in a desiccator for 30 min. Mass Spectrometry. MALDI-MS spectra and MALDI-MS/ MS spectra were acquired on an AutoFlex III from Bruker Daltonics (Billerica, MA) in positive-ion mode and LIFT mode, respectively. External calibration was performed using a custom peptide and protein mixture, with a calibration spot added at the nearest nongelatin area from the section. Laser raster was set to 75 μm along both x-axis and y-axis. These dimensions are the maximum resolution the AutoFlex III can achieve without overlapping adjacent laser spots. Acquired spectra were processed with FlexAnalysis 3.0 and FlexImage 2.1 (Bruker, Billerica, MA). LCMS/MS spectra were acquired on an Orbitrap Velos from Thermo Scientific (West Palm Beach, FL) coupled to a nanoflow LC system (NanoAcquity, Waters Corp, Boston, MA). 8796
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Figure 2. (a) Fluorescent 40 ,6-diamidino-2-phenylindole (DAPI) staining of a spheroid showing the locations of living cells; (b) fluorescent positive proliferating cell nuclear antigen (PCNA) staining showing the location of the proliferative outer ring of cells; (c) overlay of PCNA and DAPI staining; (d) colorimetric PCNA staining, PCNA is indicated by dark brown; (e) trypan blue exclusion staining, showing the location of necrotic cells in the central core. Scale bars in parts ac are 200 and are 500 μm in parts d and e. Background subtraction for parts ac was performed with Image J.
Histology Staining. Spheroid sections were treated with histology staining. Intact spheroids were submerged in 4% trypan blue solution from Sigma-Aldrich (St. Louis, MO) for 30 min. The stained spheroids were transferred to PBS solution. The spheroids were washed with PBS for 30 s. The GAS step was repeated to obtain optical images of the sections with an inverted microscope at 10 magnification. Colorimetric Staining. Proliferating cells were identified using a monoclonal mouse-antihuman proliferating cell nuclear antigen (PCNA) antibody (BioGenex, San Ramon, CA) as the primary antibody during overnight incubation, followed by HRPconjugated rabbit-anti mouse IgG (Serotec, Raliegh, NC) for 1 h. The slides were then developed in 3,3-diaminobenzidine (DAB) as the substrate (Biomeda, Foster City, CA) for 1 min and a hematoxylin QS counterstain was applied (Vector Laboratories, Burlingame, CA). The slides were then coverslipped with permanent mounting medium, and optical images of the sections were acquired with an inverted microscope at 10 magnification. Fluorescent Staining. Proliferating cells were detected using a mouse antihuman PCNA monoclonal antibody (BioGenex) as the primary antibody for 1 h. The slides were then treated with Image-iT (Invitrogen, Eugene, OR) for 30 min and incubated in goat antimouse IgG conjugated to Alexa-Fluor 488 (Invitrogen) for 30 min at room temperature. 40 ,6-Diamidino-2-phenylindole (DAPI) (Invitrogen) was then applied to the slides as a counterstain, and the slides were coverslipped using ProLong Gold (Invitrogen). The images were captured with a Nikon Eclipse 90i fluorescent microscope (Melville, NY) at 10 magnification with DAPI or FITC filters. The exposure time was set to 500 ms.
’ RESULTS AND DISCUSSION This study was performed with spheroids grown with the colon carcinoma cell line, HCT 116. (Figure 1ad). Similar to a poorly vascularized tumor, the center of the spheroid will often grow under hypoxic conditions and undergo cell death, resulting in a necrotic core. In contrast, the outer rim of cells has ample access to nutrients and oxygen and proliferates rapidly. The differences in growth can be visualized by classical methods (Figure 2). Positive staining for proliferating cell nuclear antigen (PCNA) (Figure 2b,d) indicates that cell proliferation is primarily localized in the outer regions of the sections. Trypan blue staining, which labels nonviable or dead cells, is concentrated in the central core (Figure 2e). These classical immunological methods provide valuable information about specific, known targets. However, to achieve multiplex determination of unknown species, we employed a mass spectrometric-based approach. The aim of this study was to develop a MALDI-IMS approach to examine protein distributions in 3D cultures. To do so, we have adapted several of the standard MALDI-IMS protocols used to prepare tissue sections as well as developed new approaches for manipulation of the 3D cultures. 3D cell cultures are heterogeneous structures of varying sizes, depending on the cell type used. The colon carcinoma spheroids used in this study typically grow to 1 mm diameter. To facilitate manipulation of the cultures and prevent deformation during sectioning, we embedded our samples in gelatin. Gelatin embedding of small samples for MALDI-IMS has previously been described.39 Embedding the spheroids in a solid 8797
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Figure 3. Spatial distribution of analytes in a single HCT 116 spheroid. Ion intensity maps and mass spectra are displayed for four species. While m/z 12 828 is located predominately in the central necrotic core, the distribution of other species is more widespread throughout the structure.
medium stabilizes the sample and provides a smooth cutting surface for sectioning. Traditionally, OCT (optimal cutting temperature polymer) is used when preparing tissue sections for routine histological work; OCT is not recommended for mass spectrometric applications due to the background signal of OCT in MALDI-MS analysis.40 Gelatin is a good alternative39 because matrix crystal formation is negligible on gelatin, resulting in limited background matrix signal. The concentration of the gelatin solution determines the cutting results, with more concentrated solutions providing minimal deformation of the sample. Careful sample preparation is critical for a successful MALDIIMS experiment. Isopropanol solutions are used to wash away lipids from the cell membrane and extract the analytes from the sample. However, the longer wash times can lead to the dissociation of cells. To preserve maximum sample integrity, we modified the traditional washing procedure by decreasing the washing time to less than 10 s.
The thickness of the section will affect the mass spectrometry results as well. The overall peak intensity and the total number of observable peaks increase as the sections become thinner,38 but there is a risk of losing cells with rinsing steps, resulting in large hollow areas in the section. As a compromise, we cut the samples 1012 μm in thickness so as to keep the culture geometry intact while achieving the maximal mass spectral signal. Also, desiccating the slides after the matrix application improves ionization of the analytes. Using MALDI-IMS, we can map the spatial distribution of specific analytes in 3D cultures. Working with colon carcinoma spheroids, we generated multiple slices of individual spheroids and obtained mass spectral images. Representative spectra and an ion density map from an HCT 116 spheroid slice are shown in Figure 3. Most m/z values are detected in most cells in the section (m/z 10 086, 10 495, and 11 290). However, m/z 12 828 is localized to a centralized region in the spheroid, corresponding to the necrotic core. 8798
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Figure 4. MALDI-MS ion intensity maps generated from 6 consecutive 10 μm slices of a single HCT 116 spheroid. The slices are in order of vertical position within the structure, with 60 μm vertical spacing between each two slices. The signal for m/z 12 828 is present only in the central regions of the spheroid, as it is only detected in the center of the structure.
In order to demonstrate that analyte distribution is reproducible throughout one spheroid, we imaged several m/z values in one culture. We generated 16 consecutive 10 μm slices from a single spheroid, with 60 μm of vertical spacing between each of the slices. Ion density maps for five m/z values are shown for six of the central slices in Figure 4. As well as generating spatial maps of analyte distribution, we identified proteins in the spheroids. To do so, we employed a combined strategy of direct fragmentation via MALDI-TOFTOF and by nanoflow-LCMS/MS of spheroid cell lysates and compared detected proteins against the molecular weights observed in MS images. Our identification strategy is as follows.
Spheroids were grown and homogenized, and lysates were fractionated with 1D-gel electrophoresis. On the basis of m/z values generated from the tissue sections, bands with matching molecular weights were extracted from the gel and subjected to tryptic digestion. For example, the gel bands at roughly 12 kDa were extracted and digested, as many species were observed in that mass range in the IMS ion maps. Digested samples were spotted on MALDI plates and injected for LCMS/MS analysis. Masses of identified proteins are then compared against m/z values detected in the IMS sections. Cytochrome C and histone H4 were both identified with this approach (Supplemental Table 1 in the Supporting Information). A sample MS/MS spectrum 8799
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Figure 5. MALDI-MS/MS spectrum generated from the extract of an in-gel tryptic digestion of an HCT 116 spheroid lysate. Fragmentation of the m/z 1168.66 parent ion provides the peptide sequence TGPNLHGLFGR, from cytochrome C.
generated via MALDI-TOF-TOF corresponding to a peptide from cytochrome C is shown in Figure 5, and an MS/MS spectrum from a histone H4 peptide is shown in Supplemental Figure 1 in the Supporting Information. Cytochrome C is a heme protein associated with the electron transport chain of the mitochondria. It has numerous functions, most notably as a regulator of both respiration and apoptosis.41 Given its link to apoptosis, we would have expected cytochrome C to be more concentrated in the inner necrotic core of the spheroids, but it was detected throughout the structure. Similarly, histone H4 was uniformly detected in the spheroid slices. As histone H4 is one of the core protein components of chromatin,42 it is not surprising that it is widely expressed. With a globular domain and an N-terminal tail, it is a structural component of the nucleosome in eukaryotic cells and is subject to covalent modification, which acts in diverse biological processes including gene regulation, DNA repair, chromosome condensation, and spermatogenesis.43,44
’ CONCLUSIONS We have developed a promising method to examine the distribution of biological molecules in the 3D culture systems. Our methodology does not suffer the same limitations of immunological work, where a priori knowledge of the analytes is required. In order to image these structures, we adapted gelatin-assisted sectioning to allow manipulation of the cultures. We have demonstrated that the chemical composition across the spheroids can be probed with MALDI-IMS and that individual species can be identified. Future work will focus on improvements to the spatial resolution for the cultures. Also, in addition to mapping the spatial distribution of peptides and proteins, we will expand the technology to examine distributions of lipids and other biological species. The expansion of MALDI-IMS to manipulatable 3D cell culture systems is an exciting advance that will have applications ranging from basic research to clinical discoveries.
’ ASSOCIATED CONTENT
bS
Supporting Information. Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected]. Phone: 574-631-0583. Fax: 574631-6652.
’ ACKNOWLEDGMENT The authors wish to acknowledge the assistance of Dr. William C. Boggess and Dr. Michelle V. Joyce of the Notre Dame Mass Spectrometry & Proteomics Facility. We would also like to thank Sarah Chapman of the Notre Dame Integrated Imaging Facility. Funding was provided by the University of Notre Dame and the 2011 Starter Grant from the Society for Analytical Chemists of Pittsburgh. ’ REFERENCES (1) Debnath, J.; Brugge, J. S. Nat. Rev. Cancer 2005, 5 (9), 675– 688. (2) D’Souza-Schorey, C.; Chavrier, P. Nat. Rev. Mol. Cell. Biol. 2006, 7 (5), 347–358. (3) Birgersdotter, A.; Sandberg, R.; Ernberg, I. Semin. Cancer Biol. 2005, 15 (5), 405–412. (4) Gaedtke, L.; Thoenes, L.; Culmsee, C.; Mayer, B.; Wagner, E. J. Proteome Res. 2007, 6 (11), 4111–4118. (5) Kunz-Schughart, L. A.; Freyer, J. P.; Hofstaedter, F.; Ebner, R. J. Biomol. Screen. 2004, 9 (4), 273–285. (6) El Haj, A. J.; Cartmell, S. H. Proc. Inst. Mech. Eng., Part H 2010, 224 (12), 1523–1532. 8800
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