Patient samples
Clinical samples were obtained from 20 castration resistant prostate cancer patients who were recruited from November 2008 to August 2009 at the University of California San Francisco (UCSF) to participate in our study (Table
1). Twenty patients were selected based on statistical considerations for a pilot study. All patients gave informed consent under a protocol approved by the UCSF Institutional Review Board. Blood was collected in three CellSave™ tubes (Veridex). This allowed us to use one tube for an initial CTC enumeration using the published CellSearch™ method, and the two remaining tubes to be combined for IE/FACS isolation. Microdissection and DNA extraction of archived formalin-fixed paraffin-embedded (FFPE) tumor biopsy samples from two patients were done in a manner as previously described [
7]. Using H&E stained slide, the pathologist (J.S.) demarcated tissue containing at least 80% tumor. The H&E stained slide was then used as a guide to microdissect the tumor area using a surgical blade (Feather #15) on non-stained deparaffinized sections (10 sections × 5 μM thickness). In the case of a small tumor (Patient #20), manual microdissection of the selected area was performed by scraping successive tissue sections under a stereomicroscope.
Table 1
Enumeration and genomic profiling of circulating tumor cells (CTCs) from 20 castration resistant prostate cancer patients
1 | 1 | n.d. | N.A. | N.A. | N.A. |
2 | 1 | n.d. | N.A. | N.A. | N.A. |
3 | 85 | Yes | 1 | 50 | Yes |
4 | 12 | Insufficient sample | N.A. | N.A. | N.A. |
5 | 144 | Failed due to machine error | 1 | N.A. | N.A. |
6 | 6 | Insufficient sample | N.A. | N.A. | N.A. |
7 | 10 | Insufficient sample | N.A. | N.A. | N.A. |
8 | 2 | n.d. | N.A. | N.A. | N.A. |
9 | 590 | Yes | 1 | 20 × 2 | Yes (duplicate) |
10 | 151 | Yes | 4 | 20 | Yes |
11 | 9 | Yes | 2 | 20 | Yes |
12 | 2 | n.d. | N.A. | N.A. | N.A. |
13 | 55 | Yes | 1 | 20 | Yes |
14 | 590 | Yes | 2 | 20 | Yes |
15 | 42 | Yes | 2 | 33 | WGA failed |
16 | 0 | n.d. | N.A. | N.A. | N.A. |
17* | 54 | Yes | 1 | 20 | Yes |
18 | 33 | Yes | 1 | 20 | Yes |
19 | 34 | Yes | 2 | 20 | Hybridization failed |
20* | 12 | Yes | 1 | 18 | Yes |
Enumeration of CTCs
To increase the likelihood of isolating CTCs for molecular analysis, we first performed CTC enumeration using the automated CellSearch™ assay to identify patients with > 5 CTCs per 7.5 mL. We then isolated CTCs by IE/FACS (see below) from the remaining blood sample in these patients. Cell counts via CellSearch™ correlated closely with cell counts via IE/FACS (Park, unpublished data). In initial studies, we observed that whole genome amplification of DNA from CTCs was less efficient when CTCs were isolated after 48 h post-phlebotomy. Therefore, enumeration was performed within 24 h and isolation of CTCs via IE/FACS within 48 h after blood draw.
Cell isolation via IE/FACS
For immunomagnetic enrichment, one-half volume of Cell Buffer (Immunicon) was added to a blood sample. Immunomagnetic particles coated with EpCAM (MJ37) mAb (50 μg/10 mL of indirect conjugate or 60 μg/10 mL for the direct conjugate) and EpCAM (EBA-1) mAb conjugated to phycoerythrin (PE) (400 μL at 5 μg/mL per 10 mL of sample) were added to the sample. The sample was incubated at room temperature for 15 min, mixing at half-way point and again when incubation was completed. The sample was placed in a magnetic separator (Immunicon) at room temp for 15 min, and then aspirated to remove unbound blood cells. The bound cells were eluted in 2 mL of Cell Buffer, moved to a 12 × 75 mm tube and subjected to a second round of magnetic separation for 5 min. The fluid in the tube was aspirated and the remaining cells were resuspended in 150 μL of Cell Buffer. Twenty μL of a solution containing 2 μg of a proprietary (BD Bioscience) nucleic acid dye, and 0.1 μg of a leukocyte-specific CD45 (2D1) mAb conjugated to peridinin-chlorophyll-protein-Cy5.5 were added to resuspended cells. The sample was incubated in the dark for 15 min and then sorted using FACS Aria (BD Biosciences). CTCs were defined as nucleated, EpCAM-positive, and CD45-negative while leukocytes were defined as CD45-positive, EpCAM-negative and nucleated. Cells were sorted into 10 μL of TE (10 mM Tris-HCl 1 mM EDTA pH 8.0) in a 500 μL PCR tube. All conjugated antibodies and reagents for FACS were obtained from BD Biosciences unless otherwise noted. During FACS analysis, we sorted a range of 18-50 CTCs. Table
1 shows the number of "input cells" used for WGA, and the number of replicate analyses performed for each subject. Due to the small numbers of CTCs isolated from patients' blood, it was not feasible to perform FACS analysis after sorting. Also, since blood samples were collected in CellSave™ (Veridex) tubes [
6,
8], CTCs were thereby rendered nonviable and fixed to maintain antigenicity prior to immunomagnetic enrichment and FACS sorting. Complete FACS plots from two representative patients are shown in Additional file
1: Figure S1.
Optimization of whole genome amplification (WGA)
ACGH protocols typically use between 100 ng and 1 μg of specimen DNA in the labeling reaction, equivalent to ~20,000-200,000 cells, which does not permit reliable analysis of important samples containing small numbers of tumor cells such as CTCs in blood. Analyzing these samples for copy number aberration requires a high-fidelity whole genome amplification (WGA) step. To select the best strategy for WGA of limiting amounts of input DNA, we tested four WGA protocols: multiple displacement amplification [
9], random prime amplification [
10], ligation-mediated-PCR [
11], and the GenomePlex
® WGA4 [
12,
13]. We compared aCGH results generated from diluted BT474 breast cancer cell line genomic DNA (equivalent to ~10,000 to 10 cells) to non-limiting unamplifed BT474 DNA (600 ng). We found GenomePlex
® WGA4 to be most reliable and reproducible (data not shown). We also did not observe artifacts or bias via BAC aCGH analysis. Other groups have also demonstrated the reproducibility and reliability of the GenomePlex
® WGA protocol [
12,
14].
Whole genome amplification (WGA)
To prevent any loss of genomic material, we avoided the use of column purification of genomic DNA. Instead, whole cell lysis of isolated cells and WGA of genomic DNA were performed in the same PCR tube the cells were sorted into. Genomic DNA in the whole cell lysate served as template for WGA which was performed using the GenomePlex® Single Cell WGA4 Kit (Sigma-Aldrich) following the manufacturer's protocol. Briefly, 1 μL lysis and proteinase K digestion solution was added to cells in 10 μL of TE. The DNA was then fragmented and libraries were prepared. Amplification was performed by adding 7.5 μL of 10 × Amplification Master Mix, 48.5 μL of nuclease-free water and 5 μL WGA DNA polymerase. Samples were amplified using an initial denaturation of 95°C for 3 min followed by 25 cycles, each consisting of a denaturation step at 94°C for 30s and an annealing/extension step at 65°C for 5 min. Amplified DNA was purified using the QIAQuick PCR Purification Kit (Qiagen). DNA concentration was determined by a NanoDrop™ spectrophotometer and stored at -20°C. The average yield after WGA of ~20 CTCs is ~10 μg. Early tests via aCGH analysis of samples with WGA product yield less than < 5 μg revealed non-specific amplification. Therefore samples with low yield (< 5 μg) were excluded from further downstream assays.
Array CGH analysis
In initial studies we performed aCGH analysis using oligonucleotide microarrays (Affymetrix 50 K SNP array), but observed significant experimental variability and background noise with our WGA4 amplified DNA samples. Similarly, studies by Fuhrmann et al [
15] and Nowak et al [
14] showed that bacterial artificial chromosome (BAC) arrays outperformed oligonucleotide microarrays in detecting small gains or losses [
15] and showed far fewer outliers and less technical noise [
14]. So, we performed our aCGH analysis using a bacterial artificial chromosome (BAC) array (Version 3.2) containing 2,464 clones printed in triplicate [
16]. The BAC arrays are available at the Helen Diller Family Comprehensive Cancer Center Array Core.
The random prime amplification method [
16] was used to enzymatically label approximately 600 ng of test DNA (WGA product) or unamplified normal female genomic (Promega) reference DNA with Cyanine 3-dCTP or Cyanine 5-dCTP (Amersham), respectively. Labeled test and reference DNA were co-hybridized to a BAC array. The arrays were imaged using a custom built CCD array imaging system [
17] and analyzed using the UCSF Spot Program to calculate Cy3/Cy5 ratios. The UCSF Spot Program is a fully automated system for microarray imaging that locates both subarray grids and individual spots [
18]. Ratios are then computed based on explicit segmentation of each spot. The resultant log2 ratios were analyzed by UCSF Sproc to yield the mean and standard deviation of the log2 ratios over replicate spots. The UCSF Sproc is a companion program to UCSF Spot that maps each spot on the array to a specific clone and chromosome position, and averages over replicate spots in order to output a final ratio value for each clone on the array. Clones with a standard deviation of the log2 ratios of the triplicate spots greater than 0.2 for were excluded from analysis.
To determine copy number alterations, microarray data (sproc files) was uploaded and analyzed using Nexus 5.1 software (Biodiscovery). Data was subjected to rank segmentation, a proprietary (Biodiscovery) variation of the circular binary segmentation (CBS) [
19]. The thresholds of log2 ratio values for gains and losses were 0.2 and -0.2 respectively; the thresholds for high copy number gains and homozygous deletions were 0.6 and -0.6 respectively. Copy number gains or losses were considered recurrent if present in > 50% of the CTC samples from 9 patients.
Since sex-mismatch hybridization was performed, i.e. male test (tumor) versus female (normal) reference, we analyzed copy number alterations on chromosome X by using a feature available in the Nexus 5.1 software for automatic threshold adjustment on the X chromosome. Theoretically, the intensity of a normal male test sample on the X chromosome is 1/2 that of the normal female reference (i.e. 1 versus 2 copies of chromosome X). If the same threshold used for autosomal chromosomes was applied on X, the segmentation result will show "one copy number loss" But when corrected for gender, the segmentation analysis will no longer pick up the "copy number loss on X" because the adjusted threshold takes into account that the male sample will have 1/2 the intensity on the X chromosome compared to that of the female reference. Therefore, the automatic threshold adjustment feature allows the segmentation analysis to take into account the sex-mismatch hybridization so only copy number changes on the X chromosome that deviated from the expected normal ratio was reported. Chromosome Y was excluded in the analysis.
Concordance between aCGH profiles was measured using weighted Pearson correlation. A weighted Pearson measure was used because, in aCGH comparisons, unweighted Pearson correlation is driven by the number of ratios that differ from "normal." Since the observations in this case are the clones, one can use the spatial relationship of the clones to estimate the weights. Based on this idea, the weight of a clone was estimated by considering its deviation and that of its adjacent clone from "normal" The formula used for calculating weighted correlation coefficient (corr) is given below:
where x and y are the log2 ratio values of the two samples under consideration; w is the weight vector; ; i = 1,2,...,n-1; j = i, i + 1; n = total number of clones; and x
ref
and y
ref
are the medians of the n clones for samples x and y, respectively. Only autosomal clones were considered. Correlation coefficients falling in the intervals 0 to < 0.20, 0.20 to < 0.40, 0.40 to < 0.60, 0.60 to < 0.80 and 0.80 to 1 were said to be un-, lowly-, fairly-, moderately- and highly- correlated, respectively.