Exon capture and bulk segregant analysis: rapid discovery of causative mutations using high-throughput sequencing

Florencia Del Viso
†
Dipankan Bhattacharya
Yong Kong
Michael J Gilchrist
Mustafa K Khokha
Exon capture and bulk segregant analysis: rapid discovery of causative mutations using high-throughput sequencing
M E T H O D O L O G Y A R T I C L E Open Access
Background: Exome sequencing has transformed human genetic analysis and may do the same for other vertebrate model systems. However, a major challenge is sifting through the large number of sequence variants to identify the causative mutation for a given phenotype. In models like Xenopus tropicalis, an incomplete and occasionally incorrect genome assembly compounds this problem. To facilitate cloning of X. tropicalis mutants identified in forward genetic screens, we sought to combine bulk segregant analysis and exome sequencing into a single step. Results: Here we report the first use of exon capture sequencing to identify mutations in a non-mammalian, vertebrate model. We demonstrate that bulk segregant analysis coupled with exon capture sequencing is not only able to identify causative mutations but can also generate linkage information, facilitate the assembly of scaffolds, identify misassembles, and discover thousands of SNPs for fine mapping.Conclusion: Exon capture sequencing and bulk segregant analysis is a rapid, inexpensive method to clone mutants identified in forward genetic screens. With sufficient meioses, this method can be generalized to any model system with a genome assembly, polished or unpolished, and in the latter case, it also provides many critical genomic resources.

Background

High throughput sequencing (HTS) has revolutionized our ability to analyze genomes for mutations that cause disease phenotypes. Whole genome sequencing (WGS) can identify causative mutations in smaller invertebrate genomes such as D. melanogaster [1] and C. elegans [2] and also larger vertebrate genomes such as mouse and zebrafish [3][4][5]. However, most disease causing mutations are in exons, and WGS of vertebrate genomes remains relatively expensive compared to exome sequencing. Therefore, for disease mutation discovery, exome sequencing is high-yield at a minimum cost.

To perform exome sequencing, exons are captured using arrays [6][7][8] or in solution [9] with specifically designed probes. Since the exome represents <5% of genomes, a fraction of an Illumina HiSeq lane can generate sufficient coverage depth for confident variation detection. Much more sequencing is necessary for WGS. Exome sequencing has successfully led to discovery of mutations in polished genomes such as human and mouse [10][11][12][13][14][15][16][17].

However, exon capture HTS is limited by the quality of the genome assembly and annotation. For example, in X. tropicalis, a frog genetic model system, many of the exons have been identified, yet at least 5% of the genome, including exonic sequence, is in gaps in the latest genome assembly (v7.1) [18]. These difficulties also affect the analysis of zebrafish and other emerging model systems. In addition, even in polished genomes, a major challenge in the analysis of exomes is to identify a causative mutation amongst the large array of natural sequence variations present in the genome. In forward genetic screens, heavily mutagenized genomes have additional changes, further complicating discovery of causative mutations.

Using X. tropicalis, we aimed to circumvent these difficulties by combining exon capture HTS with bulk segregant analysis (BSA). In BSA, carriers of a recessive phenotype are crossed to produce wildtype and mutant progeny. These progeny are sorted based on their phenotype into two pools (or bulks), mutant and wildtype. Genetic markers (e.g. heterozygous SNPs, microsatellites, or RFLPs) are then assayed on the bulk DNA. If the bulks contain a sufficient number of embryos (i.e. meioses), then unlinked genetic markers will be detected as heterozygous in both pools. On the other hand, genetic markers linked to the causative mutation will appear homozygous in mutant pools but will remain heterozygous in wildtype pools. This loss of heterozygosity (LOH) in mutant pools indicates linkage and can be used to identify candidate loci. By combining data from BSA and exon capture sequencing, we can sift through the large array of sequence variations to identify a region containing the mutant gene.

In polished genomes such as mouse, exon capture/BSA can be adapted to narrow candidate mutations in a single step at much lower costs than WGS.

Xenopus is an extraordinarily valuable system for biomedical research, but forward genetic analysis only became effective with the introduction of the diploid X. tropicalis as a model [19][20][21][22]. Employing forward genetic screens, numerous mutants have been identified [23][24][25], but discovering the underlying causative mutations is difficult due to the relative paucity of genetic markers and limitations of the long-range genome assembly. Despite these difficulties, some causative mutations have been characterized [26][27][28]. To facilitate the discovery of mutations in X. tropicalis, we sought to incorporate meiotic data into our exon capture HTS by adding BSA into the sequencing step. We show that exon capture HTS with BSA facilitates local genome assembly, genetic marker identification, and causative mutation discovery. Using this method, we were able to identify candidate genes for two X. tropicalis mutants in just a few weeks. We then verified the identity of the mutated gene using either mRNA overexpression to rescue the phenotype, or morpholino knockdown to phenocopy. In one case, gaps in the genome and local misassembly would have made mutation identification extremely difficult without exon capture/BSA. This method produces genomic resources inexpensively and can quickly identify candidate mutations amongst many sequence variations.

Results

In a forward genetic screen in X. tropicalis, we identified two mutants, ruby and grinch that follow simple Mendelian inheritance, show similar phenotypes, but fall into different complementation groups ( Figure 1). Both mutants appear wildtype until stage 38-39 when edema starts to appear around the heart, steadily worsens, and finally causes death by stage 46-48 (Figure 1a,b). The etiology of the edema is unclear and could be due to cardiac, lymphatic or renal defects [29][30][31][32][33][34]. To determine the causative genes, we developed a HTS approach combining exon capture with BSA. First, we developed an exon capture array containing coding exons identified in the published X. tropicalis v4.1 genome [35]. Because approximately 5-10% of the genome is missing from this assembly [18,35], we also identified coding exons from available EST clusters and full-length mRNA sequences to augment the array.

We then captured and sequenced coding exons and flanking intronic sequence from DNA of pooled embryos, in order to exploit BSA. Hybrid carriers from our mutagenized strain (N) and mapcross strain (ICB or PopA) were crossed to generate mutant and wildtype embryos. Two pools of these embryos were collected: one pool had only phenotypically mutant (MUT) embryos and the other had wildtype (WT) embryos ( Figure 2a). We performed exon capture using our custom arrays and, using the Illumina platform, sequenced each pool individually (see Methods). We first analyzed the WT pool for heterozygosity, which identified SNPs in these populations. In mutant pools, we examined these same genomic positions for LOH suggesting linkage to the causative mutation (Figure 2a and Methods). Our analysis identified over 30,000 putative SNPs that were common across all mutant and WT pools, with a subset of SNPs becoming homozygous in the mutant pools. For the v4.1 genome assembly, we calculated a homozygosity ratio by dividing the number of homozygous SNPs to heterozygous SNPs per scaffold. We focused on scaffolds that had at least 20 SNPs per scaffold to avoid bias from falsely called SNPs (Figure 2b-e). For grinch, these excluded scaffolds (with <20 SNPs) represent less than 0.7% of the genome, and for ruby, this represents less than 0.9% of the genome. Although this introduces the risk of missing smaller scaffolds that are linked to the mutation, this filter helped clarify and prioritize our analysis. When we had completed this analysis, the v7.1 genome assembly became available so we repeated the analysis on this assembly. The v7.1 genome assembly employed a preliminary X. tropicalis meiotic map as well as synteny to other genomes to create chromosomesized super-scaffolds [18]. In this case, for all scaffolds larger than 0.5 Mb, we divided the scaffold into 0.5 Mb intervals and calculated the homozygosity ratio for these smaller intervals, again excluding intervals that had less than 20 SNPs from further analysis. This analysis identified scaffolds or genomic intervals with high homozygosity ratios indicating linkage to the causative mutation in ruby and grinch mutants (Figure 2b-e).

Cloning ruby phenotype

Prior to this analysis, we had no previous mapping information for ruby. Exon capture/BSA from mutant ruby embryos found six v4.1 scaffolds with high LOH signal ( Figure 2b) and a striking signal in scaffold 3 in the v7.1 genome (Figure 2c). Using our newly identified SNPs, we sought to both validate our SNP discovery and narrow down the interval carrying the causative mutation. For fine mapping purposes, we focused on SNPs that led to Restriction Fragment Length Polymorphisms (RFLPs) and found that 46/51 SNPs (90%) tested had the expected polymorphism. We then showed that three of the scaffolds from v4.1 (30,34,143) were tightly linked to the ruby locus, thus effectively assembling the scaffolds at this locus ( Figure 3a). Analyzing the v7.1 assembly, we confirmed that scaffold 3 contains the ruby locus and identified a misassembly of a portion of scaffold 7 (Figure 3a). Although we did not test markers on scaffolds 40 and 363 (v4.1) (Figure 2b), in the v7.1 genome, both scaffolds are incorporated into scaffold 3 and therefore are likely linked to the mutation. Scaffolds 1911 (v4.1) and 218 (v7.1) may be linked to the ruby locus, but were not tested since we mapped ruby to a 220 kb or 0.3 cM interval on scaffold 3 (v7.1) using 650 mutant embryos. This interval contains only 7 genes (Figure 3a), and analysis of exon capture sequence revealed that three of these genes, il1b, psd4, and pax8, had sequence variants leading to amino acid changes or splice site changes. Il1b had one amino acid change but was outside the mapped interval (markers 1791-2011). psd4 had 3 amino acid changes that were all conservative. More importantly, the expression pattern of psd4 did not support a role in ruby, as there appears to be little expression   prior to the onset of the phenotype (Additional file 1: Figure S1).

Pax8 had a mutation in the acceptor splice site of intron 2, causing a mis-splice and frameshift, eliminating the entire paired-box domain ( Figure 3b). We confirmed this splice site mutation by RT-PCR and Sanger sequencing (Figure 3b cDNA). Pax8 has previously been shown to be a principal and early regulator of pronephric development in Xenopus [36]. Furthermore, mutant embryos had a reduced expression of pax8 (Additional file 1: Figure S2) and defects in pronephric development as shown by pax2 ( Figure 3c) and several other pronephric markers [37] (Additional file 1: Figure S2). This is consistent with the edema phenotype, especially because the onset of the edema is shortly after the pronephros becomes functional [38].

To demonstrate that pax8 is the causative gene for the ruby phenotype, we used an antisense translation blocking morpholino (MO) to knockdown pax8. Injection of pax8 MO at the one cell stage caused a clear impairment of pronephric development in 59% of the embryos (Figure 3d) phenocopying ruby mutants. In Xenopus, we can target either the left or right pronephros by injecting one cell at the two-cell stage. When targeted to one side, pax8 knockdown causes impairment in pronephric development in 54.7% of the embryos on the injected side ( Figure 3d and Additional file 1: Figure S3). Taken together, these results indicate that the phenotype in ruby mutants is caused by a pax8 mutation that leads to a truncated protein and disrupts pronephric patterning at early stages of development.

Cloning grinch phenotype Similar to ruby, grinch mutants also develop significant ventral edema, but unlike ruby, grinch mutants have a ciliary defect that can be detected by reduced cilia-driven flow on the epidermis of mutants (Figure 4c, Additional file 2 and Additional file 3). Scanning Electron Microscopy (SEM) also demonstrates fewer and abnormal cilia in multi-ciliated epidermal cells (Additional file 1: Figure S4).

However, mapping the grinch mutation has been difficult from the start. grinch is a background mutation in X. tropicalis, identified in numerous unrelated animals c a V7 .1  3  3  3  3  3  3  3  3  3  3  3  3  7  7 Recombinants    from different labs. They all cause the same edema phenotype and based on complementation testing are at the same loci. This and the fact that the phenotype follows simple Mendelian inheritance suggest a mutation at a single gene locus, with the possibility of more than one mutant allele. Furthermore, preliminary mapping linked the grinch mutation to a 10 cM gap in the meiotic map on chromosome 10 (Figure 4a) [39]. Scaffolds spanning the interval were unknown in the v4.1 assembly, making traditional positional cloning difficult.

We employed exon capture with BSA to identify linked scaffolds. We identified nine scaffolds in v4.1 that appeared linked to this locus (Figure 2d). Aligning our exome sequences with v7.1 scaffolds, we demonstrated linkage to chromosome 10, consistent with our previous mapping. We also found linkage to scaffold 29 and a portion of chromosome 7, which indicates a misassembly (Figure 2e). Of note, the homozygosity signal in the grinch mutant pools was lower than for ruby (see below).

We next sought to validate these scaffolds as linked to the grinch locus by meiotic mapping on a panel of mutant embryos. This again highlights the power of exon capture/BSA sequencing. With thousands of newly identified SNPs, some of which were RFLPs, we used meiotic mapping to create our own local assembly and fill the gap containing the grinch locus (Figure 4a). We successfully amplified 14/17 SNPs/RFLPs and 12 of them (86%) had the expected polymorphism. Fine mapping revealed that the interval containing the mutation is at the ends of scaffold 148 and 304 (v4.1). Both scaffolds share the ccdc40 gene suggesting that either these two scaffolds are overlapping or are adjacent with a local gene duplication. In the v7.1 genome, ccdc40 is triplicated on three different scaffolds, chromosome 3, scaffold 29, and scaffold 608 (Figure 4a). In addition, one other gene, ribosomal protein L38, fell within our genetic interval. The other v4.1 scaffolds (176, 282, 317, 796, 973 and 1714) indicated by our exon capture/BSA analysis (Figure 2d) are all assembled into scaffold 10 in v7.1. Therefore, although not tested by fine mapping, they are all likely linked to the mutation.

ccdc40 in X. tropicalis is an 18 exon, 3033 bp transcript. Mutations in CCDC40 were recently found in patients with Primary Ciliary Dyskinesia [40,41]. SEM and Transmission Electron Microscopy (TEM) images of grinch cilia show similar microtubule displacement as well as shorter and fewer cilia consistent with the human disease [40] (Additional file 1: Figure S4). To assess alterations in the transcript, we analyzed pools of mutant or wt embryos and performed RT-PCR using gene specific primers within the 5' and 3' UTR regions. We then cloned and sequenced the PCR products. Overall, 20/21 clones from mutants have open reading frames that lead to truncated proteins. The remaining clone has a deletion of three consecutive amino acids (Figure 4b and Additional file 1: Figure S5). In the WT pool, which includes WT embryos as well as heterozygote carriers, 17/19 of the clones are WT. Of the two others, one has a premature stop codon and the other an internal deletion, which are both found in the mutant transcripts.

It is interesting to note that we found heterozygous SNPs in the mutant transcripts, suggesting two different mutant alleles (Additional file 1: Figure S5). This was also confirmed in exon capture sequence and may explain the relatively low homozygosity ratios seen in Figure 2d,e (see Discussion).

Within the 21 mutant transcripts cloned (Figure 4b), we found several different mutations, most of which show features of aberrant splicing leading to frameshifts and premature stop codons (Additional file 1: Figure S5). 17/21 of the clones (1-3, 6,7, 10-21) suggest a splicing defect between exons 2 and 3, but neither genomic sequencing nor the exon capture data elucidated any obvious mutations in the splice acceptor/donor sites flanking or within intron 2. grinch clones 4 and 5 (also 10,11,13,14,[16][17][18][19] retain intron 9 although again, we detect no mutations in the genomic or exon capture sequences (Additional file 1: Figure S5b). Clones 8 and 9 have a deletion of 88 bp within exon 9, whether this is due to aberrant splicing is uncertain. We consider the 1 bp insertion in clone 9 to be a PCR and sequencing artifact since it occurs after a string of "A" nucleotides and has not been detected in any other clones (Additional file 1: Figure 5b). Nevertheless all the mutant transcripts for ccdc40 are abnormal compared to wildtype.

To confirm ccdc40, we attempted to rescue the ciliadriven epidermal fluid flow using mRNA microinjection. We injected WT ccdc40 mRNA along with a GFP tracer into one cell at the two-cell stage embryos. We assayed for cilia-driven flow over the surface of the embryo using colored beads and then genotyped the embryos. ccdc40 mRNA injection rescued the severe reduction in epidermal flow in grinch mutants only on the injected side in 51/55 genotyped mutants (Figure 4c and Additional file 4 and Additional file 5).

Unfortunately, identifying the exact nucleotides mutated in grinch has not been possible despite exon capture or genomic sequencing. It is not clear if the mutant variants are due to more than one allele or a single allele that causes multiple abnormalities in splicing. However, taken together, mapping, transcript sequencing, imaging of cilia, and rescue are all consistent with ccdc40 mutation(s) causing the grinch phenotype. Given the complexity of the locus, exon capture/BSA was essential to identify this gene.

Discussion

Exon capture sequencing has transformed genetic analysis in humans and has demonstrated utility in mice.

However, to date, exon capture HTS has been scarcely reported in other genetic models [42,43] despite the significant need to facilitate cloning of mutants identified in forward genetic screens. Very recently, WGS with BSA has identified causative mutations in both zebrafish and mouse validating HTS and BSA as an effective method [4,5]. For smaller genomes, such as yeast, worms, flies or Arabidopsis WGS is probably cost efficient, therefore there is no need to pre-capture the exome. However, in model systems with large genomes, currently the cost of WGS remains relatively expensive compared to exome sequencing, especially for routine use in the large numbers of mutants. In addition, to maximize the power of BSA, depth of sequencing is useful to accurately identify SNPs especially given the challenges of unambiguously mapping short read sequences and sequence errors. In our mapping intervals, we validated~90% of the SNPs identified by exon capture/BSA. Finally, exome sequencing is high-yield since most disease mutations are in exons [44].

Despite the advantages of exome sequencing, a number of barriers exist including unpolished genome assemblies, partial genome annotation, incomplete or poor characterization of variations in strains, and incomplete inbreeding of strains. As a consequence, deciphering causative mutations from the sea of variants within an exome can be a significant challenge. There are also limits to exon capture sequence regardless of genome quality or model organism. Exon capture sequencing only identifies exons and nearby flanking sequence so causative mutations deep in introns, promoters, or enhancers can be missed. Also, insertions, deletions, and splice variants may be missed by exon capture sequence due to limitations in alignment software and short read sequencing. In each of these cases, linkage information can ameliorate these problems.

The identification of the causative mutation in ruby was rapid and the significant advantage of exon capture/ BSA sequencing was that many variants could be eliminated, thousands of genetic markers identified, and a relatively narrow interval analyzed for a causative mutation within the exon capture sequence. No preliminary mapping is necessary and the meiotic interval can be narrowed based on available meioses.

In the case of grinch, positional cloning was difficult. The linked interval was not present in the meiotic map and misassembled in both versions of the genome. Exon capture/BSA identified a large number of markers and scaffolds within the gap that allowed us to assemble the grinch locus.

However, we observed some heterozygosity at the grinch locus, suggesting more than one mutant allele. An additional allele could have been introduced since grinch was identified in numerous unrelated animals from different labs. They all cause the same edema phenotype and based on complementation testing are at the same loci. Of note, the homozygosity ratios are lower for grinch compared to ruby. The presence of another mutant allele could explain the lower homozygosity signals since each allele may be associated with different SNPs and therefore appear heterozygous. Alternatively, the mutant transcripts could be due to a single allele causing multiple abnormalities in splicing.

Regardless, all mutant transcripts had either an internal deletion or a frameshift leading to a premature stop codon. Alignment with other species shows that the C-terminus of the protein is most conserved, and we hypothesize that these truncated proteins will be nonfunctional. Furthermore, microinjection of the WT ccdc40 mRNA into mutant embryos was sufficient to rescue cilia-driven epidermal flow. Thus, despite the complexity at the grinch locus, which would have made traditional positional cloning very difficult, using exon capture/BSA, we were able to identify ccdc40 mutations as causative of the grinch phenotype.

Additionally, because exon capture/BSA is independent of the genome assembly, it can detect misassemblies. In fact, WGS/BSA experiments have identified unexpected linkage across disparate parts of the genome [4,5]. The cause was not further investigated in these studies so genome misassembly or inaccurate SNP calling remain a possibility. In our study, we found evidence of misassembly in our exon capture/BSA data for both ruby and grinch and validated these misassemblies by meiotic mapping. Once identified by exon capture/BSA, we can use meiotic mapping to reassemble the genome and continue fine mapping.

Conclusions

Based on our study, we propose the following method to rapidly clone causative mutations in model systems with large genomes, such as Xenopus, zebrafish, and mice. First, an exon capture array needs to be generated using exons from a draft genome assembly and/or transcripts from ESTs, mRNA full-length sequencing, or transcriptome studies using next generation sequencers. Second, exon capture sequencing of a wildtype pool is necessary if SNPs are not well characterized in the model system. As more individuals in a strain are sequenced, wildtype sequencing becomes less essential. Third, exon capture sequencing in a mutant pool is done to identify regions of LOH. Exon capture sequence can then be analyzed for deleterious mutations in those regions. However, in more challenging cases where the LOH interval is large or complex, we recommend validating linkage by additional fine mapping to narrow the meiotic interval further or reassemble the locus. Lastly, once a candidate gene is identified, an independent validation is necessary such as a second allele, MO phenocopy, or rescue.

For model systems with large and unpolished genomes, exon capture HTS with BSA can be critical for identifying causative mutations because so many genomic resources can be obtained inexpensively. The cost of HTS is continuously decreasing and by using multiplexing techniques like barcoding, a single Illumina sequencing lane can generate sufficient sequence at considerable depth for multiple mutants. This represents a considerable cost savings compared to WGS. Furthermore, exon capture with BSA identifies a host of SNPs, facilitates local genome assembly, and discovers candidate mutations. This can be done very quickly (weeks) compared to the laborious methods such as chromosome walking across gaps.

For model systems with polished genomes, bulk segregant analysis and HTS can rapidly identify candidate genes within a narrow locus all in a single sequencing step. Finally, developing methods to rapidly and inexpensively clone mutants strongly encourages additional forward genetic screens to discover phenotypes and gene mutations that will yield fascinating new insights.

Methods

Frog husbandry

X. tropicalis were housed and cared for in our aquatics facility according to established protocols that were approved by our local IACUC.

Mutant identification

We identified both grinch and ruby in a forward genetic screen using gamma-ray irradiation as a mutagen. We immediately recognized that grinch was in the background of our N strain since multiple, even unmutagenized animals generated the same grinch phenotype [24]. Other labs also identified a grinch phenotype (Lyle Zimmerman, Robert Grainger, personal communication), which based on complementation testing were mutations at the same loci and at the time thought to be identical alleles.

ruby was identified from a single mutagenized founder in the N strain. Although the ruby phenotype was similar to grinch, complementation testing revealed that they were at different loci. ruby and grinch animals were crossed to our mapcross strains, ICB or PopA respectively for analysis.

DNA extraction for exon capture

Either 50 mutant or wildtype tadpoles at stage 42 were collected in 15 ml of lysis buffer (20 mM Tris pH 7.5-8.0, 100 mM NaCl, 20 mM EDTA, 1% SDS) supplemented with 200 mcg/ml proteinase K. Tube was rotated end over end at 55°C overnight. A phenol extraction was followed by a phenol:chloroform extraction, and then the DNA was precipitated with isopropanol, washed with 70% ethanol, and resuspended in TE buffer. DNA was processed for exon capture sequencing according to established protocols [10].

Exon capture and sequencing analysis

We generated an exon capture array by identifying all exons in gene models present in the X. tropicalis v4.1 genome [35]. We also analyzed full-length mRNA sequences and EST clusters to see if they mapped to the genome. Unmapped sequences (due to gaps in the genome) were also added to the exon capture array. Roche-Nimblegen technical support generated probes against these targets (gene models and unmapped transcripts) using their in-house proprietary software and then manufactured the arrays.

For ruby, we generated 1 lane of 100 bp paired-end Illumina GAIIx sequencing data for each wildtype (WT) and mutant (MUT) pool generating 36.1 M pairs of reads for WT and 34.0 M for MUT. For grinch, we generated 2 lanes of 100 bp paired-end Illumina GAIIx data for each pool with 68.2 M pairs of reads for WT and 67.2 M for MUT. The Illumina reads were first trimmed based on their quality scores using Btrim [45]. We used a cutoff of 25 for average quality scores within a moving window of 5 bp. A minimum acceptable read length is 25 bp. Other parameters for Btrim were set at defaults. After the quality trimming, 96% of reads were kept for ruby WT and 89% for MUT. For grinch, the trimming passing rates were 91% and 93% for WT and MUT respectively. The trimmed reads were mapped to the reference genomes (v4.1 [accession number AAMC00000000] and v7.1 [accession number AAMC02000000] available at www.xenbase.org) using Burrows-Wheeler Aligner with default parameters [46]. The mapping results were converted into bam format first and then pileup format using samtools [47].

Using the two pileup files from WT and MUT as inputs, we created a script to analyze the alleles and their frequencies along each position of the genome with mapped reads. At each position, a minimum of 10 mapped reads in both WT and MUT pools were required for analysis. We identified a putative SNP if (i) the major allele frequency (MAF) in reads of WT is in the range of 50% and 75%; or (ii) WT major allele differs from that of MUT.

For all putative SNPs, we counted the number of homozygous SNPs (nh) in the MUT pools where MAF ≥ 90%. Then the homozygosity ratio (r) of homozygous SNPs in a region is calculated as r = nh/nt, where nt is the total number of SNPs in the same region. For reference genome v4.1, we used each scaffold as a region. For reference genome v7.1, since the scaffolds are much longer, non-overlap sliding windows with a size of 0.5 Mb was used for a region. Detailed protocols and scripts for mapping sequence data and calling SNPs is available at our website (tropicalis.yale.edu/geneticResources/ geneticResources_main.html), and SNP data is available per request.

Mapping RFLP markers were designed in regions with LOH. All the SNPs in these regions were evaluated with WAT-CUT (http://watcut.uwaterloo.ca/watcut/watcut/template. php) to generate primers flanking SNPs that create or abolish a restriction site (RFLP markers). Primers were designed to amplify at least 100-250 bp flanking a SNP so we could resolve genotypes on a simple 4% agarose gel. A list of all the primers used is described in Additional file 1: Table S1. PCR with each pair of primers was performed in 96-well plates using genomic DNA from a large panel of individual mutant embryos and WT embryos as controls. PCR conditions were as follows: 94°C for 2 min, followed by 39 cycles at 94°C for 10 s, 58°C for 30 s and 72°C for 30 s. Final extension was at 72°C for 5 min. PCR products were digested with the corresponding enzyme either 4 h or overnight at 37 º C and run in 4% 50:50 regular agarose (American Bioanalytical): NuSeive agarose (LONZA) gels.

Preliminary mapping of grinch was done using primers designed around Simple Sequence Length Polymorphisms (SSLPs) as identified by the meiotic map. PCR conditions were the same as above. Primers 148E4, 148 M1, s304-875 K, s304-219 K, 560E1, and 560B2 were designed around SSLPs with a florescent 5'-m13 tail on the Forward primer [48]. PCR was done in 2 steps. Step 1: Using 5'-m13-F primer and R-primer; PCR conditions were 94°C for 2 min, followed by 34 cycles at 94°C for 10 s, 58°C for 30 s and 72°C for 30 s, and final extension for 5 min at 72°C. Step 2: Using NED, PET, VIC, or 6-FAM-m13-F primer and original R-primer and 1/50× concentration of PCR product from Step 1; PCR conditions were 94°C for 2 min, followed by 25 cycles at 94°C for 10 s, 52°C for 30 s and 72°C for 30 s. Final extension for 5 min at 72°C. PCR products were analyzed using fragment analysis at Yale's DNA sequencing facility.

Whole mount in situ hybridization (WMISH)

We obtained clones from Open Biosystems (Thermo Scientific). The DNA templates were in vitro transcribed with T7 High Yield RNA Synthesis Kit (E2040S) from New England Biolabs to synthesize digoxigeninlabeled antisense probes. Pronephric markers clones were obtained from Open Biosystems: atp1a1, IMAGE: 6988026; slc7a8, IMAGE: 53828775: slc5a9, IMAGE: 5308256; smp-30, IMAGE: 6999181; hnf1-β, TNeu141k07; cdh16, TNeu004i09. Whole mount in situ hybridization was performed according to the standard protocol with minor modifications [49,50]. We stained embryos using BM purple at room temperature or 4°C depending on each probe. If embryos needed to be genotyped after WMISH, fixation after staining was done with 4% paraformaldehyde instead of Bouin's fixative to preserve DNA. After bleaching and equilibration in 100% glycerol, embryos were examined and photographed.

In vitro fertilization and microinjections

In vitro fertilization and microinjection were done as previously described [51] and protocols available on our website (http://tropicalis.yale.edu). Embryos were injected either at one or at two cell stage with 0.5 or 1 ng of MO solution or at 2 cell stage with up to 500 pg ccdc40 mRNA. Ccdc40 mRNA was co-injected with 100 pg GFP mRNA. After injections, embryos were left in 1/9X MR + 3% Ficoll for 1 hr and then transferred to 1/9X MR supplemented with 50 μg/ml of gentamycin. Embryos were raised at 22-27°C until the appropriate stage for fixation or evaluating phenotype. Pax8 translation blocking MO was obtained from Genetools, LLC. The sequence of the MO was: 5' ATGCTGCTGTTGGGCATCTTCCTCC 3'. MO was co-injected with Alexa 488 dye (Invitrogen) as a lineage tracer.

RT-PCR and cloning strategy

RNA was extracted from WT or from mutant embryos using Trizol solution according to the manufacturer's instructions (Invitrogen). A 1.5 μg aliquot of RNA was used to perform RT-PCR with SuperScript III Reverse Transcriptase (Invitrogen) using a gene specific primer located in the 3'UTR (UTR3-1R, 2R or 3R, Additional file 1: TableS 2) of ccdc40 transcript according to the manufacturer's instructions. PCR was performed using oligonucleotides in the 5'UTR or at beginning of the transcript (Additional file 1: Table S2). PCR was done with Phusion High-Fidelity DNA Polymerase (NEB) and conditions were as follows: 98°C for 2 min, followed by 39 cycles at 98°C for 15 s, 62°C for 30 s and 72°C for 3.5 min. Final extension was at 72°C for 5 min. Samples were then incubated with Taq polymerase for 30 min at 72°C to incorporate A overhangs for TOPO TA cloning (Invitrogen) and sequenced with several primers (Additional file 1: Table S2).

Epidermal cilia-driven flow videos

A 6 cm petri dish was coated with 1% agarose. A small wedge shaped hole was cut into the agarose to position embryos with dorsal side up. 1 μL of 1/10× solution of 5 μm diameter polystyrene beads (Bangs Laboratories) were injected dorsally. Movies were captured using Canon EOS 5D Mark II DSLR camera.

Figure 1
1ruby and grinch mutant phenotypes. ruby (a) and grinch (b) mutant phenotypes start with pericardial edema at stage (st.) 39. At st. 45, the edema worsens and is lethal. Wildtype (WT) embryos from the same cross are shown for comparison. All embryos are lateral views with dorsal to the top and anterior to the left.

Figure 2
2BSA and Exon Capture HTS. (a) Schematic of Exon Capture HTS with BSA. Two pools of embryos are sequenced, one mutant and one WT from a mapcross. The WT pool is sequenced and SNPs identified based on allele counts where the major allele fraction (MAF) is approximately 50%. These positions are examined in the mutant pools for LOH. (b,d) Graph of homozygosity ratio to scaffold number using genome v4.1 for ruby (b) and grinch (d). (c,e) same as b,d but version 7.1 of the genome. In (b-e), scaffolds or scaffold intervals with <20 SNPs have been excluded. The peaks with the highest homozygosity ratios are labeled.

Figure 3
3Mapping of ruby mutation. (a) Mapping interval derived from exon capture and BSA. The number of recombinants for each RFLP marker, the genetic distance and the location of the marker in v4.1 and v7.1 genome assemblies are shown. The 220 kb interval contains 7 genes: tmem150a, c2orf68, usp39, sftpb, pax8, psd4 and a portion of il1b. Genes in red have mutations causing amino acid or splice site changes based on the exon capture data. (b) Genomic, cDNA and protein sequences of WT and mutant pax8. The mutation of two contiguous nucleotides (nt) in intron 2 (arrowhead) shifts the acceptor splice site 1 nt, causing the inclusion of an extra G in the transcript (cDNA) (arrowhead). The protein sequence shows a change in frame and shortly after a premature STOP codon (*) abrogating the entire paired box domain. (c) Pax2 expression in wt and mutant ruby embryos by Whole Mount in situ hybridization showing clear impairment of pronephric development (arrowhead). (d) Pax8 morpholino phenocopies ruby phenotype. Pax8 morpholino was injected at one or two-cell stage (single cell injected). Embryos were fixed at st. 36-37 and pax2 expression was used to assess pronephric development. The embryo shown for two-cell stage injection was injected on the right side (based on fluorescent dye tracer [not shown]). Arrowheads indicate abnormal pronephros. Embryos in (c,d) are lateral views with dorsal to the top and anterior to the left (in left columns) or right (in right columns).

Figure 4
4Mapping the grinch mutation. (a) Local genome assembly of the grinch locus using exon capture/BSA. The blue region marks the 10 cM gap in the meiotic map. The number of recombinants for each marker, calculated genetic distance and the location of the markers in v7.1 and v4.1 genome assemblies are shown. The 400 kb interval contains 2 genes: ccdc40 on the ends of scaffold 148 and 304 (v4.1) and ribosomal protein L38. (b) Schematic diagram of the protein sequences of the 21 mutant clones. Red regions indicate deletions, green box indicates insertions, and dashed lines indicate frameshifts. Number of bp inserted or deleted and translated amino acid (a.a.) length for each clone is indicated. See Additional file 1: Figure S5 for complete mutant protein and cDNA sequence. (c) Cilia-driven epidermal flow study using red microbeads. Arrows follow a particular bead's trajectory through time. In left column, WT embryo shows equal flow on both the left and right sides. In middle column, grinch mutants show no flow on either side. In right column, mutants injected with WT ccdc40 mRNA and GFP tracer (insert) show rescue of flow on injected side. See Additional files 1-4. Embryos in (c) are dorsal views with anterior to the right.
http://www.biomedcentral.com/1471-2164/13/649
del Viso et al. BMC Genomics 2012, 13:649
AcknowledgementsWe thank Sarah Kubek and Michael Slocum for frog husbandry and Engin Deniz for help processing movies. We thank Paul Guillod for helping with sequence analysis. We thank Lyle Zimmerman and Richard Harland for helpful discussions and feedback. FDV was supported by a James Hudson Brown-Alexander Brown Coxe Postdoctoral Fellowship. DB was supported by a UC Berkeley Undergraduate Research Apprenticeship award, an Amgen scholarship, NIH GM42341 to R. Harland, and an NIH MSTP TG T32GM07205 grant. This work was supported by the Medical Research Council (Programme number U117597137) to MJG and the NIH/NIDCR (R01DE018824 and R01DE018825 as well as an ARRA supplement to R01DE018824) to MKK.Competing interestsThe authors declare that they have no competing interests.Authors' contributions FDV, DB and MKK conceived and designed the experiments and wrote the manuscript. FDV, DB and MKK carried out the wet laboratory experiments. YK analyzed the HTS. MJG and MKK designed the exon capture array. FDV and DB contributed equally to this project. All the authors read and approved the final manuscript.
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Submit your next manuscript to BioMed Central and take full advantage of: • Convenient online submission • Thorough peer review • No space constraints or color figure charges • Immediate publication on acceptance • Inclusion in PubMed, CAS, Scopus and Google Scholar • Research which is freely available for redistribution. Del Viso, BMC Genomics. 13649Exon capture and bulk segregant analysis: rapid discovery of causative mutations using high-throughput sequencing. Submit your manuscript at www.biomedcentral.com/submitCite this article as: del Viso et al.: Exon capture and bulk segregant analysis: rapid discovery of causative mutations using high-throughput sequencing. BMC Genomics 2012 13:649. Submit your next manuscript to BioMed Central and take full advantage of: • Convenient online submission • Thorough peer review • No space constraints or color figure charges • Immediate publication on acceptance • Inclusion in PubMed, CAS, Scopus and Google Scholar • Research which is freely available for redistribution Submit your manuscript at www.biomedcentral.com/submit


Dendrimer-entrapped gold nanoparticles as potential CT contrast agents for blood pool imaging

Han Wang
Linfeng Zheng
†
Rui Guo
Chen Peng
Mingwu Shen
Xiangyang Shi
Guixiang Zhang
Dendrimer-entrapped gold nanoparticles as potential CT contrast agents for blood pool imaging
N A N O I D E A Open Access
The purpose of this study was to evaluate dendrimer-entrapped gold nanoparticles [Au DENPs] as a molecular imaging [MI]  probe for computed tomography [CT]. Au DENPs were prepared by complexing AuCl 4 ions with amine-terminated generation 5 poly(amidoamine) [G5.NH 2 ] dendrimers. Resulting particles were sized using transmission electron microscopy. Serial dilutions (0.001 to 0.1 M) of either Au DENPs or iohexol were scanned by CT in vitro. Based on these results, Au DENPs were injected into mice, either subcutaneously (10 μL, 0.007 to 0.02 M) or intravenously (300 μL, 0.2 M), after which the mice were imaged by micro-CT or a standard mammography unit. Au DENPs prepared using G5.NH 2 dendrimers as templates are quite uniform and have a size range of 2 to 4 nm. At Au concentrations above 0.01 M, the CT value of Au DENPs was higher than that of iohexol. A 10-μL subcutaneous dose of Au DENPs with [Au] ≥ 0.009 M could be detected by micro-CT. The vascular system could be imaged 5 and 20 min after injection of Au DENPs into the tail vein, and the urinary system could be imaged after 60 min. At comparable time points, the vascular system could not be imaged using iohexol, and the urinary system was imaged only indistinctly. Findings from this study suggested that Au DENPs prepared using G5.NH 2 dendrimers as templates have good X-ray attenuation and a substantial circulation time. As their abundant surface amine groups have the ability to bind to a range of biological molecules, Au DENPs have the potential to be a useful MI probe for CT.

Introduction

Molecular imaging [MI] combines conventional imaging technologies with MI probes, which are designed to detect aspects of biochemistry and cell biology that underlie disease progression and treatment response [1][2][3][4][5]. MI includes optical imaging, nuclear-based imaging (both positron-emission tomography and single photon emission tomography), and magnetic resonance imaging. Due to the difficulty of designing suitable contrast agents and probes, the use of X-ray computed tomography [CT] in MI has been limited. However, CT affords better spatial and density resolutions than other imaging modalities. These advantages become particularly apparent when CT is used to diagnose diseases in the thorax, such as lung cancer. There thus exists an urgent need to enhance the capabilities of CT by developing suitable MI probes.

Gold nanoparticles [AuNPs] have seen increasing use recently in cancer imaging and treatment [6][7][8][9][10][11][12][13][14][15][16][17][18] as they offer several advantages over conventional, iodine-based X-ray or CT contrast agents. First, because of its higher atomic number and electron density, gold has a higher X-ray absorption coefficient than iodine, endowing it in principle with a greater ability to enhance contrast on CT [19]. AuNPs also appear to be biocompatible [20,21]. It is relatively easy to modify the surface of AuNPs with functional groups such as targeting molecules or specific biomarkers, endowing the resulting particles with characteristics favorable for a range of MI applications [22][23][24]. Finally, proper treatment of AuNPs can increase their circulation time in the cardiovascular system [CVS] by allowing them to avoid removal by the reticuloendothelial system [RES] [22,25,26]. This is particularly advantageous when treating tumors, whose combination of leaky vasculature and poor lymphatic drainage can result in what is known as the enhanced permeation and retention [EPR] effect. Extended circulation times can exploit the EPR effect to enhance transport of AuNPs to the tumor site, while in parallel, the particles' bound targeting molecules increase the rate of endocytotic uptake [22,25].

Dendrimers are a class of highly branched, synthetic, and spherical macromolecules comprising a wide array of types, chemical structures, and functional groups [27]. Two types are commercially available: the poly (amidoamine) [PAMAM] and poly(propylene imine) dendrimers. Both types can be synthesized to different generations, each generation increasing proportionally in size and molecular weight. Thus, for example, generation 4 [G4] PAMAM dendrimers are approximately twice the size of generation 3 [G3]. These compounds have several advantages for clinical use. Not only are they highly soluble in aqueous solutions, but also their size can be precisely controlled. In addition, the terminal amine groups can easily be acetylated to shield their positive potential, thereby avoiding nonspecific binding and toxicity [28,29]. Dendrimers also possess a hollow interior that can be used to trap AuNPs as well as a substantial number of available surface amino groups that can be modified by a range of targeting molecules [30][31][32]. Dendrimers thus have a considerable potential as a nanoplatform to create multifunctional, dendrimerentrapped gold nanoparticles [Au DENPs] [33,34], which have the additional benefit of being stable not only in water, phosphate-buffered saline [PBS], and cell culture medium, but also at different temperatures and pH conditions [35]. In this study, we synthesized and characterized Au DENPs and performed a preliminary evaluation of their ability to attenuate X-rays in vitro and their in vivo use as a MI probe for CT.

Materials and methods

Synthesis and characterization of Au DENPs

Generation 5 PAMAM [G5.NH 2 ] dendrimers with a polydispersity index less than 1.08 were purchased from Dendritech (Midland, MI, USA). All other chemicals were obtained from Aldrich (St. Louis, MO, USA) and used as received. The water used in all the experiments was purified using a Milli-Q Plus 185 water purification system (Millipore, Bedford, MA, USA) with a resistivity higher than 18 MΩ cm. Regenerated cellulose dialysis membranes (molecular weight cutoff, 10, 000) were acquired from Fisher (Waltham, MA, USA).

Au DENPs were synthesized using previously reported methods [30,35,36] with minor variations. Briefly, the particles were prepared using sodium borohydride reduction chemistry, with gold salt/dendrimer molar ratios of 51  [36]. To determine the size distribution of the {(Au 0 ) 200 -G5.NH 2 } DENPs, a 1 mg/mL aqueous solution of each sample was dropped onto a carbon-coated copper grid and allowed to air-dry. The grids were then viewed by transmission electron microscopy [TEM] using a JEOL 2010F analytical electron microscope (JEOL, Tokyo, Japan) operating at 200 kV. For each sample, 300 Au DENPs were randomly selected for size analysis, which was performed in parallel by three investigators. The size-distribution histogram was produced using ImageJ software (http://rsb. info.nih.gov/ij/download.html).

In vitro CT imaging and CT value measurement

Fifteen serial dilutions of either {(Au 0 ) 200 -G5.NH 2 } DENPs or iohexol (Omnipaque ® , 300 mg iodine per mL, GE Healthcare, Milwaukee, WI, USA), ranging from 0.001 to 0.1 M of [Au] or [I], were prepared in 1.5-mL microcentrifuge tubes and placed in a self-designed scanning holder. The tubes were then scanned using a 64-row multidetector CT system (LightSpeed VCT, GE Medical Systems, Milwaukee, WI, USA) with the following parameters: tube voltage, 120 kV; tube current, 50 mA; slice thickness, 0.625 mm; slice space, 0; scan field of view, 25 cm; display field of view, 6 cm; matrix, 512 × 512. Each concentration of either {(Au 0 ) 200 -G5.NH 2 } DENPs or iohexol was scanned three times, with a 24-h interval between any two scans.

Images were reconstructed, and CT values measured, using a GE imaging workstation (Advantage Workstation 4.3, GE Medical Systems, Milwaukee, WI, USA). Images were reconstructed in the axial plane, after which a 20-mm 2 circle was laid over the center of each image to define the region of interest for the measurement of CT value. CT values were calculated based on three scans of each sample, each performed by a different investigator, and the data were presented as mean ± standard deviation.

In vivo CT imaging

The institutional animal care committee of Shanghai Jiao Tong University approved all animal experiments. BALB/c mice (20 to 25 g, Shanghai Laboratory Animal Center) were anesthetized by intraperitoneal injection of 3% sodium pentobarbital (35 mg/kg). The mice were scanned using a micro-CT imaging system (eXplore Locus, GE Healthcare, London, Ontario, Canada) set to the following parameters: tube voltage, 80 kV; tube current, 450 μA; exposure time, 400 ms; slice thickness, 45 μm; slice space, 0; scan field of view, 45 mm × 80 mm; effective pixel size, 0.046 mm. Images were reconstructed on a micro-CT imaging workstation (GEHC microView, GE Healthcare, London, Ontario, Canada) using the following parameters: voxel, 45 μm × 45 μm × 45 μm; display field of view, 10 to 25 mm.

To test the performance of Au DENPs as a CT MI agent, a 10-μL aliquot of {(Au 0 ) 200 -G5.NH 2 } DENPs with [Au] of 0.007, 0.009, 0.01, or 0.02 M was subcutaneously injected into the back of the experimental mice while the control mice were injected with an equal volume of PBS (pH 7.4). These Au DENP concentrations were chosen on the basis of both the CT value of the soft tissue area in the uninjected mice and the previously derived CT value measurements of Au DENP solutions alone. After injection, micro-CT scans and image reconstruction were carried out as described above. Each experiment was carried out three times.

In vivo dynamic digital X-ray photography

Dynamic digital X-ray photography was carried out using a mammography system (Senographe DS, GE Medical Systems, Milwaukee, WI, USA) set to the following parameters: tube voltage, 22 kV; tube current, 8 mA; exposure time, 400 ms. The anesthetized mice were imaged before injection and then 5, 20, and 60 min after injection of either Au DENPs or iohexol. The acetylated {(Au 0 ) 51.2 -G5.NHAc} DENPs (300 μL, [Au] = 0.2 M), 300 mg/mL iohexol, or PBS were injected into the tail vein at a flow rate of 300 μL/min. Images were interpreted on a picture archiving and communication system monitor (Pathspeed, GE Medical Systems Integrated Imaging Solutions, Mt. Prospect, IL, USA) after adjustment of the optimal window settings, and then analyzed. This part of the study was performed by two investigators in consensus.  Figure 2) shows that the particles were relatively uniform in size, forming a normal distribution.

Results

Synthesis and characterization of Au DENPs

In vitro CT imaging and CT value measurement

The reconstructed CT images obtained by scanning various concentrations of either {(Au 0 ) 200 -G5.NH 2 } DENP or iohexol solutions are shown in Figure 3. CT values (in Hounsfield units [HU]) derived from these scans ( Table 1) were used to construct the concentration-CT value curves shown in Figure 4. These showed, first, that at a concentration of 0.01 M or less, X-ray attenuation by Au DENPs was slightly less than that observed with an iohexol solution containing the same concentration of iodine. However, these differences were small, within 6 HU. In contrast, as the concentration was In vivo CT imaging Figure 5 shows that {(Au 0 ) 200 -G5.NH 2 } DENPs ≥ 0.009 M could be detected by micro-CT imaging after being injected subcutaneously into the dorsum of the mice [35]. After injection, the Au DENPs tended to become distributed as a short segment in the interspace between the skin and the subcutaneous soft tissue.

Dynamic digital X-ray photography

At the 5-and 20-min time points after injection of {(Au 0 ) 51.2 -G5.NHAc} DENPs, the vascular system of the mice was visible on CT ( Figure 6I-b, 6I-c), with the heart, renal vein, main portal vein, and branches of the portal vein each clearly evident. The urinary system could be distinguished at the 60-min time point, with the ureter and urinary bladder defined clearly. In contrast, injected iohexol was unable to image the vascular system. At the 5-and 20-min time points after iohexol injection, the urinary system was imaged only vaguely, and after 60 min, only the urinary bladder was imaged. Together, these findings indicated that Au DENPs remained in the vascular system longer than iohexol and provided superior imaging enhancement.

Discussion

In this study, we compared the ability of Au DENPs and iohexol to attenuate X-rays in vivo and in vitro as well as their ability to persist in the circulation after intravenous injection. We found a normal {(Au 0 ) 200 -G5.NH 2 } DENP size distribution around 4.0 ± 0.9 nm. The CT  AuNPs hold a considerable promise as CT contrast agents for blood pool imaging because AuNPs persist longer in the circulation and exhibit a five-to seven fold higher attenuation of X-rays as compared with iodine-based agents [6,26,37]. According to the Lambert-Beer law [38], the relationship among an input X-ray flux I 0 , a tissue matrix of thickness T with linear attenuation coefficient μ m , and the transmitted flux I m is described by the formula I m = I 0 ·e -μmT . When both tissue and contrast agent are present, the flux I c that passes through a scanned section of thickness t is I 0 ·e -μmT-t ·e -μct , or I m ·e -(μc-μm)t , where μ c is the linear attenuation coefficient of the contrast agent. The difference in the signal between the surrounding matrix and the feature defined by the contrast agent can then be calculated as C = (I m -I c )/I m = 1 -e -(μc-μm)t . Thus, the difference in signal intensity induced by a contrast agent is introduced depending only on the thickness of the contrast agent and the difference in the linear attenuation coefficients of the contrast agent and the matrix. For this reason, the attenuation coefficient of a given contrast agent is one of the most important factors that determine its CT imaging efficiency.

Comparison of the concentration-versus-CT value curves of {(Au 0 ) 200 -G5.NH 2 } DENPs and iohexol indicated that increasing the molar concentration of either element led to an increase in its attenuation coefficient. This was likely due to a concentration-dependent effect caused by the change in mass ratio between water molecules and either [Au] or [I]. The CT value of Au DENPs indicated that they had the superior ability to attenuate X-rays. Together, these results indicated that Au DENPs had a significant potential for use in CT MI based on their ability to enhance contrast. To further explore the feasibility of using Au DENPs in CT MI, we used {(Au 0 ) 200 -G5.NH 2 } DENP solutions for micro-CT imaging and {(Au 0 ) 51.2 -G5.NHAc} DENPs for dynamic digital X-ray photography in vivo. At concentrations above 0.009 M, Au DENPs had a much higher attenuation coefficient     [35]. Effective MI probes must have a sufficient half-life in the vasculature as enough of the agent must be transported to the target site to allow data collection [25,39]. This is especially true when MI agents are used for tumor diagnosis. Tumor vessel walls are incomplete and fragile, containing large gaps between the endothelial cells and the basement membranes [40]. This makes tumor neo-vessels highly permeable, allowing contrast agents to diffuse freely from the vasculature into the interstitial space. However, the half-life of a nanoparticle contrast agent is also determined by its size [27,41,42]. Molecular probes that are less than 3 nm in diameter, such as G1 or G2 dendrimers, leak easily across vessel walls into the surrounding tissue. Larger particles, between 3 and 5 nm in diameter, such as G3 or G4 dendrimers, are quickly excreted through the kidney, making them potentially useful as functional renal contrast agents. Agents between 5 and 8 nm diameter, such as G5 or G6 dendrimers, are retained in the circulation and are thus best suited for use as blood pool contrast agents or MI probes. However, when the diameter exceeds 20 nm, the agents are easily taken up by the RES within the liver and spleen.

In view of the previous considerations, and as the objective of this study was to produce a probe with a maximum half-life in vivo, we used G5.NH 2 dendrimers as templates to prepare Au DENPs. The images obtained after injection of Au DENPs or iohexol indicate that Au DENPs remained in the circulation longer than iohexol.

Previous studies have shown that the entrapment of AuNPs within dendrimer templates does not influence the surface properties of the dendrimers [35]. Thus, the Au DENPs we synthesized would be expected to retain the native ability of the PAMAM dendrimer, allowing effective chemical modification with biologically active molecules [30].

Although the current results are promising, further experimental studies will be needed to ensure that these Au DENPs are effective and safe for clinical application. For example, the ability of these molecules to be modified so as to target particular organs and tissues should be evaluated, and the ability of organs and tissues to take up thus modified particles specifically measured. Before clinical application is considered, the potential toxicity must be ruled out.

This preliminary study demonstrates that Au DENPs prepared using G5.NH 2 dendrimer templates have good X-ray attenuation and a substantial circulation time in the CVS. Their potential to be biologically and chemically modified [43], combined with ongoing improvements in computer technologies and the spatial resolution of CT scanners, is likely to make CT MI with these and related agents an increasingly important tool for diagnosis and drug delivery.

Figure 1
1shows a typical TEM image of the synthesized Au DENPs prepared with a gold salt/dendrimer molar ratio of 200:1. The size of the {(Au 0 ) 200 -G5.NH 2 } DENPs was estimated to be 4.0 ± 0.9 nm. The size-distribution histogram (

Figure 1
1TEM images of {(Au 0 ) 200 -G5.NH 2 } DENPs. 0.01 M, X-ray attenuation by Au DENPs became progressively greater than that of the corresponding iohexol solution.

Figure 2
2Size-distribution histogram of {(Au 0 ) 200 -G5.NH 2 } DENPs.

Figure 3
3Axial CT images. {(Au 0 ) 200 -G5.NH 2 } DENPs (a) and iohexol (b) at a range of concentrations in 1.5-mL microcentrifuge tubes. {(Au 0 ) 200 -G5.NH 2 } DENPs exceeded that of iohexol at Au concentrations above 0.01 M. {(Au 0 ) 200 -G5.NH 2 } DENPs ([Au] ≥ 0.009 M, 10 μL) were detectable by micro-CT after subcutaneous injection. The vascular system could be imaged 5 and 20 min following the injection of {(Au 0 ) 51.2 -G5.NHAc} DENPs into the tail vein, and the urinary system could be imaged after 60 min.

Figure 4 Figure 5
45Concentration-CT value curves of {(Au 0 ) 200 -G5.NH 2 } DENPs and iohexol. Micro-CT images of the experimental mice. The mice were injected subcutaneously with 10 μL {(Au 0 ) 200 -G5.NH 2 } DENPs at [Au] of 0.007 (a), 0.009 (b), 0.01 (c), and 0.02 M (d). The white circle in (a) indicates the injection site. Arrows in the remaining panels show where Au DENPs have become distributed as a short segment in the interspace between the skin and the subcutaneous soft tissue. The mean CT values at the injection region were 31.57 (a), 41.23 (b), 48.56 (c), and 75.76 HU (d).

Figure 6
6Planar projection images after intravenous injection of {(Au 0 ) 51.2 -G5.NHAc} DENPs or iohexol. Rows I, II, and III contain images obtained after injection of the Au DENPs, iohexol, and PBS, respectively. In each row, image (a) is the pre-contrast image, (b) was taken 5 min after contrast injection, (c) at the 20-min, and (d) at 60 min after contrast injection. The following structures could be clearly distinguished 5 min after Au DENP injection (I-b): the heart (arrow), renal vein (oval arrow), main portal vein (arrow head), and branches of the portal vein (diamond arrow). At 20 min (I-c), the renal vein (oval arrow), main portal vein (arrow head), and branches of the portal vein (diamond arrow) remained distinct. Sixty minutes after Au DENP injection (I-d), the vascular system could no longer be visualized, but the urinary system, including the ureter (open arrow) and the urinary bladder (open arrow head), could be seen distinctly. After iohexol injection (II), the vascular system of the experimental mice could not be imaged. The urinary system began to be imaged 5 min after iohexol injection (II-b), and after 60 min (II-c), only the urinary bladder was defined.

Table 1
1CT values of Au DENPs and iohexol solutions.Concentration (M)
CT value (HU)

Au DENPs
Iohexol

0.001
0.9 ± 1.9
5.1 ± 2.8

0.002
4.2 ± 2.3
10.3 ± 2.8

0.003
11.6 ± 2.1
16.3 ± 1.3

0.004
14.9 ± 1.4
20.8 ± 1.5

0.005
20.1 ± 2.2
26.8 ± 1.6

0.006
23.3 ± 1.4
31.2 ± 1.4

0.007
33.7 ± 1.5
35.6 ± 2.3
0.008
35.8 ± 1.9
40.1 ± 2.2

0.009
39.2 ± 2.9
44.9 ± 2.3

0.01
42.3 ± 8.7
48.3 ± 2.2

0.02
76.3 ± 5.8
69.1 ± 3.7

0.04
159.7 ± 18.7
158.6 ± 10.2

0.06
238.1 ± 15.6
180.7 ± 12.5

0.08
325.3 ± 23.3
233.9 ± 18.2

0.1
546.7 ± 27.1
286.5 ± 16.7

CT, computed tomography; Au DENPs, dendrimer-entrapped gold
nanoparticles.

After acetylation to transfer dendrimer terminal amine groups to acetamide groups, the dendrimer tertiary amines cannot stabilize the larger Au DENPs entrapped within the dendrimers. The results obtained using both {(Au 0 ) 200 -G5.NH 2 } and {(Au 0 ) 51.2 -G5.NHAc} DENPs are comparable in terms of X-ray attenuation intensity since Au DENPs prepared using different gold salt/dendrimer molar ratios with a size range of 2 to 4 nm display similar X-ray attenuation at similar Au concentrationthan the parenchyma, allowing very low-dose amounts
of Au DENPs to be visible within the parenchyma on
the CT image. We utilized acetylated {(Au 0 ) 51.2 -G5.
NHAc} DENPs for intravenous injection because acety-
lation of the terminal amines of Au DENPs can signifi-
cantly improve their biocompatibility by avoiding the
amine-induced toxicity that can arise at high concentra-
tions [28,30,35,36]. Noting that although {(Au 0 ) 200 -G5.
NH 2 } DENPs are stable at PBS and cell culture medium
[35], acetylation of {(Au 0 ) 200 -G5.NH 2 } DENPs cannot
generate stable {(Au 0 ) 200 -G5.NHAc} DENPs. This is
because the {(Au 0 ) 200 -G5.NH 2 } DENPs have a relatively
larger size (4.0 nm) when compared with {(Au 0 ) 51.2 -G5.
NH 2 } (2.1 nm) [36].
© 2012 Wang et al; licensee Springer. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
AcknowledgementsThis work was supported by grants from the National Natural ScienceAuthors' contributions HW carried out the design of the imaging studies and participated in the in vitro and in vivo imaging studies and the manuscript drafting. LZ participated in the imaging studies and the manuscript drafting. RG carried out the design of the nanoparticle studies and participated in the synthesis and characterization of Au DENPs and the manuscript drafting. CP participated in the synthesis and characterization of Au DENPs. MS participated in the design of the nanoparticle studies. XS and GZ conceived the study and participated in its design and coordination. All authors read and approved the final manuscript.Competing interestsThe authors declare that they have no competing interests.
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. Wang, Nanoscale Research Letters. 7190Wang et al. Nanoscale Research Letters 2012, 7:190