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Determination of enriched histone modifications in non-genic portions of the human genome

Background Chromatin immunoprecipitation followed by high-throughput sequencing (ChIP-seq) has recently been used to identify the modification patterns for the methylation and acetylation of many different histone tails in genes and enhancers. Results We have extended the analysis of histone modifications to gene deserts, pericentromeres and subtelomeres. Using data from human CD4+ T cells, we have found that each of these non-genic regions has a particular profile of histone modifications that distinguish it from the other non-coding regions. Different methylation states of H4K20, H3K9 and H3K27 were found to be enriched in each region relative to the other regions. These findings indicate that non-genic regions of the genome are variable with respect to histone modification patterns, rather than being monolithic. We furthermore used consensus sequences for unassembled centromeres and telomeres to identify the significant histone modifications in these regions. Finally, we compared the modification patterns in non-genic regions to those at silent genes and genes with higher levels of expression. For all tested methylations with the exception of H3K27me3, the enrichment level of each modification state for silent genes is between that of non-genic regions and expressed genes. For H3K27me3, the highest levels are found in silent genes. Conclusion In addition to the histone modification pattern difference between euchromatin and heterochromatin regions, as is illustrated by the enrichment of H3K9me2/3 in non-genic regions while H3K9me1 is enriched at active genes; the chromatin modifications within non-genic (heterochromatin-like) regions (e.g. subtelomeres, pericentromeres and gene deserts) are also quite different.

Background
The chromatin state and transcription level of a chromosomal region has been found to be related to modifications of histones [1,2] as well as DNA [3,4]. The fundamental structural unit of chromatin is the nucleosome, which is formed by wrapping DNA around a histone octamer consisting of two copies each of four core histone proteins (H2A, H2B, H3, and H4). The tails of these histones can be modified in a variety of ways that relate to chromatin condensation and gene expression [2]. Different histone residues can be methylated, acetylated, phosphorylated or ubiquitinated to either directly change the chromatin structure or allow for the binding of specific transcription factors [5].
The acetylation and methylation states of various lysine residues have been extensively investigated. Due to the reactive nature of the amine group on the side chain of lysine, it can be acetylated or mono-, di-or tri-methylated. Enrichment for acetylated histones has been found to correlate with euchromatic and gene-coding regions [6,7]. In contrast, certain methylations are associated with activation and others with repression. The most extensively characterized activating methylation is H3K4 [8][9][10], while the H3K9 [11][12][13] and H3K27 [14][15][16] methylations are thought to lead to repression and inactivation. Recently, though, mono-methylations of H3K27 and H3K9 have been found to be enriched in active genes [17].
Using antibodies with specificity for a particular methylation state, cells have been profiled separately for mono-, di-, and tri-methylation of the same residue. These comparisons were originally performed using mass-spectroscopy and comparisons of staining patterns [18][19][20]. Since these studies relied on comparisons of staining patterns, they were only able to produce low-resolution results that could not be linked to sequence positions. More recently, the ChIP-chip [21] and ChIP-seq [17] techniques have been utilized for high-resolution sequence level profiling of histone modifications in mammalian genomes [22,17]. These techniques utilize chromatin immunoprecipitation coupled with either tiling microarray hybridization or high-throughput sequencing to determine the genomic locations of a particular modification. It is now possible to obtain resolution on the level of individual nucleosomes. Because histone modifications act on individual nucleosome and there may be modification differences between adjacent nucleosomes, this level of resolution is highly informative.
The most comprehensive profiling of histone modifications has been performed in CD4 + T cells [17,23]. The genome-wide locations at a single nucleosome level resolution of 38 different histone methylations and acetylations were profiled, including different levels of methylation of the same residue. In these studies, the modification profiles surrounding the promoters of genes with different levels of expression were examined. Basic comparisons between the different modifications were performed in the region surrounding annotated genes, but here we are reporting the results of an extension of our initial studies [17,23] to non-genic regions of the human genome.
In particular, we wanted to investigate histone modifications in three important largely non-coding regions of the genome: pericentromeres, subtelomeres, and gene deserts. These regions are mostly non-coding, but little work has been performed to characterize and differentiate them with regard to histone modifications. Using the identified genome-wide locations for histone methylations and acetylations in CD4 + T cells, we have identified the modifications that are strongly enriched in pericentromeres, subtelomeres, and intergenic regions (we have excluded all known genes in those regions). We have also used consensus repeat sequences to analyze histone modifications present in the unassembled portions of telomeres and centromeres.

Characterization of histone modifications enriched in pericentromeres, subtelomeres or gene deserts
To characterize the histone modifications that are enriched in different regions of the genome, we first obtained the locations of each type of region. Our determinations were made based upon gene annotations (see Methods and Figure 1 for details). Pericentromeres were defined for each chromosome as the region between the two genes most proximal to the centromeric gap in the genome assembly. From this region, the 10 kb nearest to each of the genes was removed since it could contain regulatory elements for the genes. Subtelomeres designations from [24] were used as a starting point. If a gene was found in a subtelomere, the gene was removed along with the 10 kb surrounding it on each side. Gene deserts were defined as regions of greater than 1 Mb between two adjacent genes, with the 10 kb proximal to each gene removed. From all three types of regions, any known Pol II binding sites [17] or DNase I hypersensitive sites [25] with 100 bp flanking regions were removed in order to exclude regions that may be related to the adjacent genes.
In each of these genomic regions, we determined the statistically significant histone modifications. First, the average number of tags for each modification in each type of region was determined. Random samples of the genome having the same size as each type of region were then taken and the number of tags for each modification was determined for the samples. By comparing the actual count to the counts in the random samples, an empirical p-value for significance could be determined (see methods for details). The results of this analysis are shown in Table 1. All of the modifications shown are significant with a q-value < 0.05, with the full data listed in Additional File 1.
The results for pericentromeres are intriguing. The enrichment of H3K9me3 is expected, as it is a known mark for constitutive heterochromatin. H4K20me3 is a known mark of repressed chromatin and therefore it would be expected to be enriched in non-genic regions of the genome. Surprisingly, it is only found strongly in pericentromeres and in no other type of non-genic region. The appearances of H4R3me2 and H3R2me2 as significant modifications in pericentromeres are likewise interesting. Methylation of H4R3 has been generally correlated with expression and active chromatin [26,27]. But, as with other histone modifications, no distinction has been made between the two methylation states of H4R3, H4R3me1 and H4R3me2. Therefore, H4R3me2 could be a signal of pericentromeres while H4R3me1 could be a signal of active genes. This is supported by the fact that H4R2me2 levels were not found to have a preferential enrichment at either active or silent promoters [17]. H3R2me2 has been found to impede the binding of effector proteins to H3K4me3 and thus to control gene expression [28]. It is possible that the significant levels of H3R2me2 in pericentromeres are important in preventing the expression of genes in those loci. These four modifica-A diagram outlining the analysis that was performed on the data Figure 1 A diagram outlining the analysis that was performed on the data. The diagram uses the example of gene deserts to describe the analysis, but the same techniques were used for pericentromeres, subtelomeres and silent, medium expression and high expression genes. tions are all strongly correlated with each other within pericentromeric regions indicating that they are likely to be found on the same nucleosomes.
For gene deserts, the di-and tri methylations of H3K9me3 were found to be significantly enriched, as would be expected [29]. Distinctly lacking, were H3K27me3 and H3K9me1. This absence could be due to the presence of H3K27me3 and H3K9me1 in silent and active genes (see below). For subtelomeres, the repressive marks H3K27me2/3 were found to be enriched along with two acetylations that are marks of activation, H2AK5ac and H3K14ac.

H4K20 and H3K9 methylation levels are varied in pericentromeres, subtelomeres and gene deserts
To further characterize the histone modifications in different non-genic regions, we compared the multiple methylation states of individual residues in each region. The number of tags identified for each methylation state of H4K20 and H3K9 in pericentromeres, subtelomeres and gene deserts were counted and the results are shown in Figure 2. For H4K20me, the differences between the types of chromatin are significant. In each region, levels of H4K20me1 are minimal and invariant. One possible reason for this is that H4K20me1 is a marker of active genes [17,30] and it is depleted in constitutive heterochromatin. The reverse is found for H4K20me3 which has been found to associate with constitutive heterochromatin [31]. In pericentromeres, the level of H4K20me3 is over 40 times the level of H4K20me1. This difference decreases in gene deserts and it is lowest in subtelomeres where the level of H4K20me3 is only double the level of H4K20me1. This indicates that H4K20me3 may be important for pericentromeres.
We next compared the three levels of methylation of H3K9 ( Figure 2B). The levels of H3K9me2/3 have been found to be enriched in non-genic regions, while H3K9me1 levels are enriched at active genes [17]. We found that for each type of non-genic region the order of enrichment levels is the same with H3K9me3> H3K9me2>H3K9me1. While the order is the same, the ratio between the methylation levels varies. In pericentromeres, the levels of H3K9me3 are almost double those of H3K9me2. This difference is greatly decreased in subtelomeres where the levels are similar and within one standard error of the mean (see error bars). In pericentromeres, the combined levels of H3K9me2 and H3K9me3 are 11 times those of H3K9me1. This is reduced to 4 times the level of H3K9me1 in subtelomeres.

Analysis of histone modifications at centromeric and telomeric repeats
Since large regions of the centromeres and telomeres of the human genome are not included in the genome assembly, they could not be accurately probed by our techniques. To probe these regions, we have used a A comparison of the enrichment of different levels of methylation at the same residue in gene deserts, pericentromeres and subtelomeres Figure 2 A comparison of the enrichment of different levels of methylation at the same residue in gene deserts, pericentromeres and subtelomeres. The mean has been plotted and the error bars show plus and minus one standard error of the mean. All of the data has been normalized to the number of counts of each modification in the genome, so the genomic average for each mark is 1.
technique similar to that previously used for studying histone modifications at repetitive regions in the mouse genome [32]. As a proxy for the actual genome sequences at repetitive locations, consensus repeat sequences are used. We have taken all of the reads for each methylation and determined the percentage of them that aligned either to telomere or centromere consensus repeat sequences.
Since the percentages of repeats were calculated and compared, no normalization for differences in antibody specificity or number of sequenced tags were required. The percentage is a self-normalizing calculation. (see Methods for details) For centromeres, the most represented modifications are H4K20me3 and H3K9me3 and the least significant modifications are H3K4me1 and H4K20me1. Both H4K20me3 and H3K9me3 are known to be associated with constitutive heterochromatin [29,33] and were also found to be significantly enriched in pericentromeres. This indicates that centromeres have heterochromatic characteristics and may require these histone modifications to prevent expression. For telomeres, the most significant modifications are H2BK5me1 and H3K4me3 and the least significant are H3K36me3 and H3K9me3. The finding of these two activating modifications to be the strongest in telomeres is interesting and is likely to be related to the recent finding of RNA polymerase II transcription at mouse [34] and human [35] telomeres. The lack of H3K9me3 also indicates that telomeres are not consistent with other non-coding regions of the human genome. In contrast to the middle of a chromosome where pericentromeres and centromeres have similar enriched histone modifications, the ends of chromosomes at subtelomeres and telomeres have different modifications. We also found that a much larger percentage of the tags matched the centromeric repeats than the telomeric repeats. The percentage of tags matching the centromeric repeats range from 0.02% to 0.8%, while the range for telomeric repeats is from 10e -4 % to 10e -3 %. This can be explained by the much larger percentage of the genome contained in centromeres than in telomeres [36,37]. Because the percentage of reads was compared rather than the raw numbers of reads, this result should not be affected by differences in sequencing depth.

Determination of modifications with statistically significant enrichment in different regions of the genome
We next decided to compare our findings for non-genic regions to the genic portions of the genome. Using comparisons to random samples of the genome (see above and methods), we determined the histone modifications statistically significant in silent genes, medium expression genes, high expression genes, and a combined set of all genes without regard to expression. The complete list of significant modifications is summarized in Table 2. All modifications listed in this table are significant at a qvalue of 0.05 and a complete list of all modifications with their mean counts and empirical p-values and q-values are in Additional File 1. When all of the genic regions without regard to expression were tested, all of the acetylations and some of the methylations were found to be significant. Interestingly, when the genes were divided into groups by expression level, different modifications were found to be significant for each level of expression. The least well-characterized groups are silent genes and nongenic regions which each only have a few modifications significantly enriched. In contrast, there are many modifications that are preferentially associated with active genes.

H3K9me2/3 and H3K9me1 have variant levels of enrichment in non-genic regions and in genes
Since the significance analysis found H3K9me2 and H3K9me3 to be enriched in non-genic regions and H3K9me1 enriched at active genes, we wanted to further investigate the relationship between H3K9 methylation state and expression level. In general, H3K9 methylation has been considered to be an indicator of inactive and silent chromatin [13,11,12], but increased levels of H3K9me1 are found consistently in the gene body of active genes [17]. Interestingly, certain active genes have also been found to have H3K9me3 [38][39][40].
To investigate the roles of mono-, di-and tri-methylated H3K9, we compared the levels of enrichment of each modification in overall non-genic regions in addition to genes with different levels of activity ( Figure 3A). It was found that the overall profiles for H3K9me2 and H3K9me3 are highly similar (Pearson correlation of 0.99). The enrichment for these two modifications was highest in non-genic silent regions and decreased as the activity of the regions increased. In contrast, a completely opposite profile was found for H3K9me1. As the activity level for genomic regions increased, the level of H3K9me1 increased greatly. These findings indicate that H3K9me is not a monolithic modification and H3K9me1 is distinct from H3K9me2/3.

The three levels of H3K4 methylation are all activating marks, but H3K4me3 is the least enriched in active genes relative to inactive genes
H3K4me has long been considered an activating mark for chromatin, but differences between H3K4me2/3 and H3K4me1 have been found [8,17,41]. To compare the enrichments of H3K4me1, H4K4me2, and H3K4me3 in broader regions, we compared the counts of each modifi-cation in regions of the genome ( Figure 3B). This figure suggests that the enrichment patterns for H3K4me1, me2, and me3 are almost congruent (Pearson correlation of 0.99) with each other indicating that all three levels of methylation of H3K4 are equally important determinants of expression level as activating marks. The only regions of the genome having a significant difference between the modifications of H3K4 are highly expressed genes. In these regions H3K4me1 and H3K4me2 are enriched at higher levels than H3K4me3 The methylation states of H3K27 each have a distinct profile Different methylation states of H3K27 are found to be significant in distinct parts of the genome ( Table 2). Because of this variation, we wanted to determine how the different modification states are related to genes with different levels of expression. We determined the number of counts for each methylation state of H3K27me3 in each type of region in the genome. The resulting plots are shown in Figure 4.
It was found that the levels of H3K27me3 are most enriched at silent genes. The levels of this modification is similarly minimal in both non-genic regions and active genes. Rather than having an enrichment level that is either correlated or anti-correlated with expression, it has A. A comparison of the profiles of the different levels of methylation of H3K9 from non-genic regions to high expression genes a parabolic profile with the peak in silent genes. This finding is especially interesting when considered in concert with the profiles of the other levels of methylation of H3K27 and their presumed functions. Each level of methylation of these two residues has a completely distinct profile. This diversity is much greater than what was found for any other histone residues including H3K4, H4K9 and H4K20.

Discussion
We have analyzed histone modification patterns in pericentromeres, subtelomeres, and gene deserts. It was found that each region has a distinct histone modification profile even though they are all traditionally considered to be constitutive heterochromatin or heterochromatin-like [42]. This is not unexpected since each of them has a distinct function. Pericentromeres are involved in the cohesion of sister chromatids during cell division [43], subtelomeres maintain chromosome stability [24,44] and gene deserts do not have a well defined function but they may contain long-range regulators of gene function [45,46] or structural regions such as the nuclear matrix attachment sites [47,48]. Each of these functions is performed by a specific set of DNA binding proteins and thus should be associated with distinct histone modifications. One possibility for the discrepancy between H3K9me3 levels in gene deserts and pericentromeres as compared to telomeres relates to their locations in the genome. Pericentromeres and gene deserts are in the middle of a chromosome between genes (even though the genes may be at a great distance) and therefore they need H3K9me3 which will interact with HP1 [49] to prevent transcription from occurring. Subtelomeres do not require such an enrichment of H3K9me3.
A further conclusion from the results in Table 1, is that the modifications found in gene deserts are a subset of the modifications that are found at pericentromeres. Both contain repressive H3K9me2/3, but pericentromeres also have H3R2me2, H4K20me3 and H4R3me2. This difference may be due to the replication associated functions of pericentromeres [50]. This is in addition to the requirement of a constitutive heterochromatin sequence at centromeres [43].
A recent study [32] looked at constitutive heterochromatic regions of the genome by aligning ChIP-seq reads to consensus sequences for repetitive elements and found that H3K9me3 and H4K20me3 were strongly enriched. We have reached a similar conclusion, but were able to separate the centromeric and telomeric regions and distinguish the modifications that are enriched in each of these regions. We found that the enrichment of H3K20me3 in non-coding regions is largely due to its high level in pericentromeres and that it is found at lower levels in subtelomeres and gene deserts.
The comparisons between non-coding regions and genes with different levels of expression have allowed us to determine the histone modifications enriched in each region. We have found that silent genes have a distinct histone modification profile that is not merely an intermediate between genes with high levels of expression and nongenic regions. Non-coding regions are characterized by high levels of H3K9me2/3 whereas silent genes are characterized by H3K27me3. This is contrary to previous studies that have grouped H3K9me2/3 and H3K27me3 as repressive modifications [29,51] without identifying significant differences between them. Our higher resolution and sharper distinction between the genomic regions allowed for this differentiation.

Conclusion
We have found that regions of the genome that do not contain actively expressed genes do not have a monolithic histone modification profile. Silent genes as well as different sub-categories of non-coding regions including gene deserts, subtelomeres and pericentromeres each have different significantly enriched histone modifications. We have also determined the histone modifications at highest levels in non-assembled telomeres and centromeres.
Profiles of different levels of methylation for H3K27 Figure 4 Profiles of different levels of methylation for H3K27. The points represent the mean value with the error bars showing the standard error. Each level of methylation of this residue has a distinct profile with the tri-methylation most highly enriched in silent genes. Even though the levels of the three types of H3K27me are indistinguishable in silent genes, the levels of H3K27me3 are clearly higher there than anywhere else. All of the data has been normalized to the number of counts of each modification in the genome, so the genomic average for each mark is 1.
Additionally, we have found that for each histone residue studied different methylation states are enriched in different parts of the genome. It is therefore an oversimplification to speak about the function of methylation of a histone residue without specifying the methylation state under discussion.

Chromatin modification data
The chromatin modification data was obtained from [23,17].

Determination of genomic regions and types of chromatin
The genic regions of the genome were the UCSC Known Genes (April 2007 release). To remove redundancy, splice variants of genes were combined and each exon was only included once in the set. Additionally, only genes that had accurate unambiguous mappings to Affymetrix U133 Plus 2.0 arrays were used. This procedure resulted in a list of 12,544 genic regions. For all of the analyses, the gene region refers to all of the sequence from the start to end of transcription.
The genes were divided into three groups based upon their expression level in CD4 + cells [52]. The high expression genes are the 1000 genes with the highest level of expression, the medium expression genes are the middle 1000 genes when the genes are sorted by expression and the silent genes are the 1000 genes with the lowest level of expression.
The gene desert regions were determined by first identifying adjacent genes in the genome with a 1 Mb break between adjacent genes. From these regions, the outer 10 kb on each end that could contain regulatory elements for the neighboring genes were excluded. To further purify the sequence, any known DNase hypersensitive sites [25] and RNA Pol II binding sites [17] were removed along with 100 bp flanking regions on each side of these sites The subtelomeric regions were taken from [53]. The original data from that manuscript http://www.wistar.org/ riethman/ contained genomic coordinates for the July 2003 (hg16) build of the human genome. These coordinates were mapped onto the Mar. 2006 (hg18) assembly using the UCSC LiftOver tool. Of the 21 Mb of sequence that were identified in hg16, 13 Mb could be accurately mapped to hg18.
The pericentromeric regions were determined based upon the centromere locations identified in the UCSC Genome Browser. The sequence between the two most proximal genes on either side of the pericentromere was obtained. From this sequence, any known DNase hypersensitive sites [25] and RNA Pol II binding sites [17] were removed along with 100 bp flanking regions on each side of these sites.
In order to refine the regions that were being used and to eliminate the number of false negative that were found, we utilized the identified nucleosome locations within the human genome [52] as a control. For a histone to be modified, there needs to be a nucleosome present, thus any region in the genome that is lacking nucleosomes by definition, must lack all histone modifications. In addition, there are regions throughout the human genome that cannot accurately be mapped accurately using the short reads [54,55]. We therefore only included those regions of the genome in which nucleosomes were found. We mapped all of the nucleosome reads from [52] to the human genome and then extended the read length to 150 bp to include the entire length of the nucleosome. This resulted in an effective genome size of 2,171,857,609 bp. The percentages of each type of region that was covered by nucleosomes and therefore analyzed are in Additional File 2.

Determination of counts in particular regions
The determination of the number of counts for a particular modification in a genome region will be outlined using the example of H3K4me3 and high expression genes. This same method was used for each modification and each type of genomic region. All of the high expression genes were extracted and divided into non-overlapping 1 kb windows. For each window, the number of H3K4me3 tags whose middle was within the window was counted. This count was then normalized by dividing it by the average number of counts for H3K4me3 in 1 kb windows in the whole genome covered by nucleosomes. Through this normalization step, the counts of different modifications were made comparable. The raw counts are affected by differences in the affinities between antibodies for different modifications and therefore have different distributions. A normalized count was determined for each window and the mean and standard error of the mean were calculated. This mean appears in the graphs as the normalized count per kb for H3K4me3 in high expression genes. The standard error of the mean is shown in the error bars.

Statistical Significance of Modifications
The average counts of each modification in all regions of a particular type were compared with random samples of the nucleosome filled portions of the genome (see above). The rank of the actual count in the distribution of the randomized counts gave an empirical p-value. Using the example of H3K4me3 in high expression genes this will be illustrated. The total count of H3K4me3 marks in the entire gene body from the TSS to the end of transcription for all high expressed genes was determined. Then, ten thousand random samples of the nucleosome-filled genome each having the same total number of bases as that of all of the high expressed genes were made. For each random sample, the total number of counts of H3K4me3 was determined. These sample counts were used to create a null distribution for the number of H3K4me3 tags that would be found in a random portion of the genome covered by nucleosomes. The actual count was then compared to the null distribution to calculate an empirical pvalue. In essence, this randomization determined how strongly H3K4me3 was enriched in high expressed genes relative to genes with lower levels of expression and noncoding regions. The p-values that were produced from this randomization were corrected for multiple testing using a false-discovery rate (FDR) technique [56,57]. All modifications that are marked as significant in a particular region of the genome are significant at a q-value of 0.05.

Centromeric and Telomeric Repeats
For centromeric and telomeric repeats, the consensus sequences in RepBase [58] were used. All of the tags for each modification were collected and compared to the specified repeat sequence using RMAP [55]. Since RMAP is designed as a program for aligning short sequencing reads to a genome, it needed to be used in a modified fashion. The "genome" that the reads were compared to was the consensus sequence for either centromere or telomere repeats. Thus, running RMAP with the entire list of H3K9me3 reads and the telomere repeat sequence, would give the number of H3K9me3 reads that mapped to the telomere repeat. This count was then divided by the total number of H3K9me3 reads to give the percentage of reads mapping to telomeres. A percentage was used in order to correct for differences between the read libraries for the different histone modifications that could be affected by many different factors including antibody specificity and the number of sequencing runs performed. If there were twice as many reads for modification A as for modification B, the percentages would still be directly comparable because modification A would have twice as many total reads along with twice as many telomere reads which would produce the same percentage or mapped reads.


extfieldsofstudy: Medicine Biology

text:
New occurrence of B chromosomes in Partamonahelleri (Friese, 1900) (Hymenoptera, Meliponini)

Cytogenetic analyses of the stingless bee Partamona helleri collected in the state of Bahia, Northeast Brazil revealed the chromosome numbers n = 18 in the haploid males and 2n = 35 in the diploid females. All karyotypes displayed one large acrocentric B chromosome, which differs from the minute B chromosomes previously described in the populations from southeastern Brazil. Giemsa staining, C-banding and DAPI/CMA3 fluorochrome staining also revealed a remarkable interpopulational divergence regarding both the regular karyotype and the B chromosomes. The B chromosomes found in the samples from Jequié, Bahia, were entirely heterochromatic, while those found in Cravolândia, Bahia, displayed a euchromatic portion at the telomeric end of the long arm. CMA 3 labeling sites varied from seven to eight between the two localities in Bahia, due to the presence of an extra GC-rich block in the karyotype of the samples from Jequié. This is the first report of a large B chromosome in P. helleri and reveals the occurrence of a geographic differentiation within this species.

Partamona is a Neotropical genus of stingless bees with an ample distribution in a wide variety of habitats going from southern Brazil to central Mexico. These habitats include rain forests, cerrado (Brazilian savanna), caatinga, and highlands. Given the existence of morphologically similar species, this group has a problematic taxonomy. Some species can only be recognized by their nesting behavior or nest entrance architecture (Pedro and Camargo, 2003).
Of the 33 species currently recognized (Pedro and Camargo, 2003), only eight, P. pearsoni (Tarelho ZVS, 1973, MSc Dissertation, University of São Paulo), P. seridoensis (Brito-Ribon et al., 1999, 2005), P. aiylae, P. vicina, P. mulata, P. nhambiquara (Brito-Ribon et al., 1999), P. peckolti (Brito et al., 2003), and P. helleri (Costa et al., 1992;Brito et al., 1997Brito et al., , 2005, were cytogenetically studied. All these species showed the regular chromosome number 2n = 34, but P. helleri showed a diploid numeric variation ranging from 2n = 34 to 2n = 38, due to the occurrence of up to four minute B chromosomes per individual (Costa et al., 1992;Brito et al., 1997Brito et al., , 2005Tosta et al., 2004). B chromosomes are extra chromosomes to the regular complement and are characterized by their dispensability, independent evolution, and non-Mendelian patterns of inheritance (Beukeboom, 1994). Their origin is a matter of recurrent debate among cytologists. Some have proposed that the appearance of the B chromosomes suggests the involvement of rearrangements in the regular chromosomes, e.g., centric fragment formation through chromosome fusions (Camacho et al., 2000). An alternative hypothesis suggests an origin through interspecific hybridization as observed in the fish Poecilia formosa (Schartl et al., 1995) and in the wasp Nasonia vitripennis (McAllister and Werren, 1997). Brito et al. (1997) distinguished two morphological types of B chromosomes occurring in P. helleri in southeastern Brazil. Although the presence of the minute B's was common in the previously studied population, the fourfold B dosage [2n = 38 chromosomes, Tosta et al. (2004)] was rare. This finding suggests the existence of a mechanism controlling or preventing the accumulation of B chromosomes in individuals.
potentially useful for analyzing the frequency, geographic distribution, transmission, or effects of the B chromosome in the adult organism. However, despite the contributions given by previous studies, the origin of the B's in P. helleri is so far still unclear.
Here, we report a new type of B chromosome in populations from the northern limits of the distribution of P. helleri in the state of Bahia, Brazil. Differences from previous descriptions found in the regular karyotype are also discussed.
Metaphases were obtained from cerebral ganglia of prepupae, following the protocol described by Imai et al. (1988). A total of 50 specimens were analyzed.
We performed C banding according to Sumner (1972), with the slight modifications proposed by Pompolo and Takahashi (1990). DAPI/CMA 3 staining was done as described by Schweizer (1980). C-banded and conventional Giemsa-stained chromosomes were analyzed and photographed with a Leica DLMS photomicroscope using HQ film. Fluorescent images were captured in a Leica DMRA2 photomicroscope using the IM50 software. In order to compare our results with previous studies, the karyotypes were arranged by decreasing order of length of euchromatic arms, and the chromosome nomenclature followed Imai (1991), with metacentric chromosomes showing pericentomeric (M cc ), centromeric (M c ), pericentromeric and telomeric (M cct ), centromeric and telomeric (M ct ), and pericentromeric and interstitial (M cci ) heterochromatic bands (see the original reference for more details).
In the present analyses, the karyotype of P. helleri showed 35 chromosomes in the females and 18 chromosomes in the males. For the Cravolândia samples, the Cbanded diploid karyotypic formula found was 2k = 28M cc + 4M c + 2M cct +1B (Figure 1d).
The novel B chromosomes found in the Bahia samples were much larger than those described in the population from southeastern Brazil (Brito et al., 1997(Brito et al., , 2005. They had an acrocentric morphology and a size comparable to the chromosomes of the regular complement. The B chromosomes observed in the Jequié samples were entirely heterochromatic, whereas those found in Cravolândia showed a euchromatic portion at the telomeric end of the long arm (Figures 1c, d). These results reveal a substantial karyotypic divergence among the populations studied so far, due to structural changes in both the regular and the B chromosomes. Brito et al. (1997) classified the minute B's found in the southeastern Brazilian populations as B1 (heterochromatic submetacentrics) and B2 (acrocentrics, with C-banding undetermined due to their small size). Partially and entirely heterochromatic B chromosomes were found in another bee species, Melipona quinquefasciata (Marla P. Rocha, personal communication).
The DAPI fluorochrome stained the heterochromatin in all centromeric and pericentromeric regions of regular chromosomes (Figures 2b, c), but the CMA 3 fluorochrome staining was variable. Specimens from Jequié showed eight CMA 3 + labeling sites, seven on the regular chromosomes and one on the B chromosome (Figure 2a). In the samples from Cravolândia, only seven CMA 3 labeling sites were observed, six on the regular chromosomes and one on the B chromosome (Figures 2c). The latter result agrees with Brito et al. (2005), who also found seven CMA 3 + markings on regular chromosomes and one on the minute B in P. helleri from southeastern Brazil. These authors also found a heteromorphism in the second chromosome pair due to a difference in the size of the heterochromatic/CMA 3 + long arm, a result not observed in the Bahia populations. Similar heterogeneous AT-and GC-rich heterochromatin blocks were observed in other meliponine species, such as Plebeia sp. and Melipona sp. (Maffei et al., 2001), and Partamona peckolti (Brito et al., 2003).
The present analysis using different techniques revealed the occurrence of a geographic differentiation between the populations of P. helleri. We observed karyotypic differences in the size, morphology, and distribution of the heterochromatin in the chromosomes of the regular complement and of the B chromosomes. Our results show that the B chromosome system of P. helleri is more complex than previously assumed. The large B chromosomes found in the Brazilian Northeast region can be an important element for future evolutionary studies of this species. In the study carried out by Tosta et al. (2007), the authors found a SCAR (Sequence Characterized Amplified Region) marker associated with the minute B chromosomes of animals from the southeastern region. This was the first attempt to use molecular data to investigate the ef-fect of B chromosomes on the individuals and showed that the approach could be effective for future population studies of this species, especially concerning the origin of the B's.
The karyotypic differences described indicate that the B chromosomes found in the present study and those previously reported in the southeastern region could have had distinct origins, probably involving rearrangements in several of the regular chromosomes. The so far undefined origins and patterns of the B chromosome inheritance can be clarified with an ample analysis of its variation and geographic distribution in this species. Other approaches, including molecular and cytogenetic data, will help to clarify the cytotaxonomy and the mechanisms involved in the karyotype evolution of P. helleri. We have already been carrying out new samplings in order to further characterize the cytogenetic and molecular diversity of this species, especially in Bahia where the large Bs were found.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine

text:
Mechanisms of the action of povidone-iodine against human and avian influenza A viruses: its effects on hemagglutination and sialidase activities

Background Influenza virus infection causes significant morbidity and mortality and has marked social and economic impacts throughout the world. The influenza surface glycoproteins, hemagglutinin (HA) and neuraminidase (NA), act cooperatively to support efficient influenza A virus replication and provide the most important targets for anti-influenza chemotherapy. In this study, povidone-iodine (PVP-I), which has a broad-spectrum microbicidal property, was examined for its inhibitory effects against influenza virus infection in MDCK cells and the mechanisms of PVP-I action on HA and NA were revealed. Results Results obtained using a novel fluorescence- and chromogenic-based plaque inhibition assay showed that 1.56 mg/ml PVP-I inhibited infections in MDCK cells of human (8 strains) and avian (5 strains) influenza A viruses, including H1N1, H3N2, H5N3 and H9N2, from 23.0–97.5%. A sialidase inhibition assay revealed that PVP-I inhibited N1, N2 and N3 neuraminidases with IC50 values of 9.5–212.1 μg/ml by a mixed-type inhibition mechanism. Receptor binding inhibition and hemagglutinin inhibition assays indicated that PVP-I affected viral hemagglutinin rather than host-specific sialic acid receptors. Conclusion Mechanisms of reduction of viral growth in MDCK cells by PVP-I involve blockade of viral attachment to cellular receptors and inhibition of viral release and spread from infected cells. Therefore, PVP-I is useful to prevent infection and limit spread of human and avian influenza viruses.

Background
Among the three types (A, B and C) of influenza viruses, A type is the most virulent, infecting various avian and mammalian species and causing human pandemics as a consequence of antigenic change (antigenic shift) in their surface glycoproteins, hemagglutinin (HA) and neuraminidase (NA) [1]. Sixteen HA and 9 NA subtypes have been recognized so far [2]. HA and NA interact with sialic acid receptors on the host cell surface, the former mediating membrane fusion that results in virus infection and the latter possessing sialidase activity that cleaves sialyl linkages between viral HA and cellular receptors to release progeny viruses and separate viruses from HA-mediated self-aggregation, allowing the virus to infect a new host cell for continuing virus replication [3].
Virus infection can be inhibited by the use of compounds that bind to viral HA [4][5][6], inhibit NA activity [7][8][9][10][11] or inhibit both HA and NA activities [12]. Two NA inhibitors, sialic acid and shikimic acid analogues, have recently been licensed for treatment of influenza A and B infections: zanamivir [13] (Relenza ® ), which is administered by inhalation, and oseltamivir phosphate [14] (Tamiflu ® ), which is administered orally as a prodrug and is converted by hepatic esterase to its active form, oseltamivir carboxylate (OC). However, influenza A and B viruses with mutations in the NA gene have developed resistance to oseltamivir and zanamivir [15,16]. The worldwide circulation of oseltamivir-resistant seasonal H1N1, highly pathogenic avian H5N1 [17,18] and the pandemic (H1N1) 2009 [19] have provided an impetus to develop new antiviral and antiseptic materials.
In the nineteenth century, povidone-iodine (PVP-I), a polyvinylpyrrolidone iodine complex, was developed and found to have a potent broad-spectrum activity against bacteria, mycobacteria, fungi, viruses and protozoa [20]. PVP-I has become widely used as an antiseptic and disinfectant. Despite long-term use, development of PVP-I resistance in microorganisms has not been reported [21,22].
PVP-I products have been found to be effective in inactivating a variety of enveloped and nonenveloped viruses, such as polio [23], herpes simplex, herpes zoster [24], and human immunodeficiency viruses [25,26]. Anti-influenza virus activity of PVP-I also has been reported recently [26][27][28]. Pretreatment of avian influenza H5N1, H5N3, H7N7 and H9N2 viruses with PVP-I products, such as solution, scrub, gargle and throat spray, in the range of 0.23-2%, reduced viral infectious titers to undetectable values in embryonated hen's eggs [27]. Both aqueous (Betaisodona ® ) and liposomal PVP-I inactivated human influenza A virus (H3N2), resulting in reduction of the virus titer by more than 4 orders of magnitude in Madin-Darby canine kidney (MDCK) cells [28]. However, the target sites and mechanisms of PVP-I action on influenza A and the other virus infections have hitherto remained unknown. In this study, we investigated mechanisms underlying PVP-I antiinfluenza activity. The apparent reduction of influenza A viral infectious titers after incubation with PVP-I products within a short period of time [26][27][28] led us to investigate two spike glycoproteins on the viral surface, HA and NA, which play essential roles in viral infection, as targets of PVP-I anti-influenza effects.

Inhibition by PVP-I of influenza A virus growth in MDCK cells
We first determined the cytotoxicity of PVP-I against MDCK cells employed as host cells of influenza viruses in this study by using a cell counting kit-8 assay. Half-maximum cytotoxic concentration of PVP-I after 24-h exposure of MDCK cells to PVP-I was 2.4 ± 0.2 mg/ml. PVP-I ranging from 0-1.56 mg/ml, which had no effect on MDCK cells, reduced virus yield in MDCK cells in a dose-dependent manner ( Figure 1B). In comparison with virus yield in the absence of the inhibitor, 1.56 mg/ml of PVP-I reduced human virus yield by 59.7-97.5% and avian virus yield by 23.0-57.4%, suggesting enhanced sensitivity towards human viruses compared to that toward avian viruses. OC, used as control, inhibited A/Memphis/1/71 (H3N2) infection by 62% and 73% at concentrations of 0.13 μM and 80 μM, respectively, whereas it inhibited A/DK/HK/ 313/78 (H5N3) infection by 20% and 37%, respectively, at the same concentrations.
Quantitative inhibition of viral HA binding to sialo-glycoconjugate receptors on the erythrocyte surface by fetuin control and PVP-I is shown in Figure 3A and summarized for PVP-I activity in Table 1. No erythrocyte hemolysis and no significant change in pH (pH of each well ranging from 6.52 to 7.20) in the assay system were observed. In general, fetuin exhibited higher inhibitory activity (ranging from 0.02 to 1.25 mg/ml) than that of PVP-I (0.2-12.5 mg/ml).

Effect of PVP-I on influenza A virus sialidase activity
In order to examine the effect of PVP-I on sialidase activity of different subtypes of influenza virus strains, the enzyme activity and K m value of each virus subtype were determined at pH 6.0 using 2'-(4-methylumbelliferyl)-α-D-Nacetylneuraminic acid (MUNA), a sensitive fluorogenic substrate without 2,3 and 2,6 linkages. Then an inhibition assay was performed using 2 enzyme units of each virus subtype and substrate concentration at its K m value. IC 50 values of OC against sialidase of different virus strains were ranged from 0.37 to 6.88 nM (data not shown).
There were marked differences in IC 50 values for PVP-I, from 9.5 to 212.1 μg/ml depending on the virus strain ( Table 1).
The kinetic mechanism by which PVP-I inhibits influenza A virus sialidase activity was investigated by determining kinetic parameters of human A/PR/8/34 (H1N1) sialidase on hydrolysis of MUNA in the absence and presence of an inhibitor. As shown in Table 2, with OC or 2-deoxy-2,3dehydro-N-acetylneuraminic acid (DANA), K m values increased, but V max did not change. In the presence of PVP-I, K m values increased and V max decreased. V max /K m ratio decreased 6-fold, 6-fold and 12-fold in the presence of 4 nM OC, 75 μg/ml PVP-I and 5 μM DANA, respectively, indicating decrease in sialidase efficiency. Lineweaver-Burk plots showed that inhibition of A/PR/8/34 sialidase activity by OC and DANA was of a competitive type, whereas that by PVP-I was of a mixed type ( Figure 4). The

Discussion
Iodine is a nonmetallic essential nutrient with a potent broad range of microbicide actions against almost all of the important health-related microorganisms, including bacteria, fungi, viruses and protozoa. Although a high content of iodine species with free molecular form (I 2 ) and hypoiodous acid (HOI) in aqueous solution has powerful microbicidal effects but can cause volatility, stinging and cytotoxicity [30][31][32]. To overcome these problems, iodine was combined with neutral carrier polymers to increase iodine solubility and to keep low the release of iodine as a solubilizing agent and to act as an iodine reservoir [30,33]. The most popular carrier in current use is povidone [32,33], which has no microbicidal activity [34]. Since povidone slowly and continuously releases free iodine into solution, these properties help to maintain antimicrobial capacity for a long period and to decrease toxicity.
By using the cell counting kit-8 assay, we found that the IC 50 cytotoxicity of MDCK cells following 24-h exposure to PVP-I was 2.4 ± 0.2 mg/ml. Based on morphological Effect of PVP-I on direct binding activity of influenza viruses to glycopolymers   [35], cell shrinkage, rounding and detachment from the surface of the culture plate after treatment with 3.1 mg/ml of PVP-I suggested that the cells were undergoing apoptosis. Therefore, we used low concentrations of PVP-I that did not cause any toxicity to host MDCK cells in order to investigate its anti-influenza virus activity.
Our results confirmed that PVP-I is a potent inhibitor of influenza virus production in MDCK cells. We indicated that PVP-I inhibits the viral replication in a dose dependent manner and is more active against human viruses (H1N1, H3N2) than avian viruses (H1N1, H5N3, H9N2). PVP-I appeared to inhibit binding of human A/Memphis/ 1/71 (H3N2) virus to specific sialoglycopolymers but not that of avian A/DK/HK/313/78 (H5N3) virus. Hemagglutination of erythrocytes induced by human viruses was inhibited by PVP-I, while hemagglutination inhibition of avian viruses required higher PVP-I concentrations. Differences in hemagglutination inhibitory activity of PVP-I against various viruses may be associated with the different structure of HA protein of each virus type. Unlike the α2,3 and α2,6 sialoconjugated protein fetuin [36], which reduces HA binding activity of both avian and human influenza viruses via competition for binding with sialyloligosaccharide receptor substrates to the viruses [37], blockage of viral HA attachment to receptor substrates by PVP-I may result from alteration of viral HA protein structure by reaction of free iodine with basic -NH groups, phenolic groups, and -SH groups of amino acid residues [30].
Although avian viruses appear to be less sensitive than human viruses to PVP-I, based on results of the erythrocyte agglutination assay, which reflects viral attachment to host cells, agglutination of avian A/DK/HK/313/78 virus (~400 HAU) was completely inhibited after a second exposure to 5 mg/ml of PVP-I. This is in agreement with the finding that titers of a highly pathogenic avian virus (H5N1) and three low pathogenic avian viruses (H5N3, H7N7 and H9N2) cultivated in embryonated eggs become undetectable by incubation with a commercial PVP-I product for 10 seconds before inoculation [27]. These results suggest that gargling with PVP-I could prevent human infection not only by human influenza viruses that bind to sialyl α2,6 Gal receptors in the upper part of human trachea but also by avian viruses that bind to sialyl α2,3 Gal receptors that exist deep in the human respiratory tract [38]. This could consequently minimize the risk of avian virus mutation, either by adaptation or reassortment, to recognize the human host predominately carrying α2,6-linked sialic acids.
PVP-I inhibited sialidase activity as a mixed-type inhibitor, indicating that free iodine is capable of binding to either free sialidase or sialidase complexed with its substrate, but iodine binding to free sialidase is more efficient than that to sialidase-substrate complex as K i was 16-fold lower than K is . This may be explained by the distribution of lysine, arginine, histidine, cysteine and tyrosine residues throughout the sequence of the NA molecule, which are reactive with iodine [30]. Although the K i value for iodine was higher than the K m value, indicating that affinity of the MUNA substrate for sialidase is higher than iodine, the activity of sialidase to hydrolyze MUNA (V max / K m = 0.12) was reduced in the presence of PVP-I (75 μg/ ml) (V max /K m = 0.02), comparable to that in the presence of OC (4 nM). The reduction in sialidase activity should result in a decrease in influenza replication.
There have been a number studies on the development of harmless carriers (such as cyclodextrin) that slowly release free iodine at a concentration retaining antimicrobial activity without a cytotoxic effect against mammalian cells for use of iodine in therapeutic applications [28,39]. Intravenous administration of iodine-lithium-α-dextrin has successfully prevented lethal infection of Staphylococcus aureus in rats [39].

Our study confirms the inhibition of avian and human influenza A virus infection by PVP-I and demonstrates
that PVP-I inhibits both viral HA binding activity and viral NA catalytic hydrolysis, mediating virus entry into host cells, and virion release and spread to a new host cell, respectively. Thus, PVP-I, for which there has been no report of resistance, is a potential agent that not only prevents viral infections but also reduces the spread of influenza viruses in epidemic and pandemic areas.

Viruses and cells
Viruses were propagated in 10-day-old embryonated chicken eggs at 34°C, and after 48 h of incubation, allantoic fluid was harvested, cleared and concentrated. The virus pellet was resuspended in cold phosphate-buffered Inhibition by PVP-I of influenza virus hemadsorption activity on guinea pig erythrocytes saline (PBS), divided into aliquots, and kept at -80°C until use. Virus titers expressed as HAU were determined (see below) before experimentation. Protein concentration was determined by using a BCA™ protein assay kit (Pierce, Rockford, IL, USA) with bovine serum albumin (BSA) as a standard. MDCK cells were cultured in Eagle's minimal essential medium (EMEM) supplemented with 5% fetal calf serum, antibiotics (penicillin-streptomycin) and glutamine at 37°C in an atmosphere of 5% CO 2 .

Cytotoxicity assay (Cell counting kit-8 assay)
Toxicity of PVP-I (Meiji Seika Kaisha, Tokyo, Japan) against MDCK cells was examined using a cell counting kit-8 (DOJINDO Laboratories, Kumamoto, Japan) to determine the numbers of viable cells using WST-8 [2-(2methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium, monosodium salt] as a substrate. In brief, confluent monolayers of MDCK cells in a 96-well plate were washed with serum-free EMEM and incubated with PVP-I at a final concentration ranging from 0-25 mg/ml for 24 h at 37°C in 5% CO 2 atmosphere. After removal of PVP-I solution, cell viability was determined by addition of kit reagent, incubation for 1 h at 37°C, and measurement of absorbance at 450 nm. PVP-I concentrations showing no cytotoxicity against MDCK cells were used in viral infection assay.

Viral infection inhibition assay
Inhibitory effects on virus development in MDCK cells were determined as shown in Figure 1A. An aliquot of virus (V) at the optimal dose (input virus causing 5 × 10 5 fluorescent forming units, which corresponds to 250 PFU/ well equivalent to a multiplicity of infection (MOI) of 0.025 PFU/cell, in the absence of an inhibitor [41]) was preincubated with an equal volume of serial dilutions of an inhibitor (I) or serum-free EMEM (100% infection) at 4°C for 1 h (Pretreatment step). A 100-μl aliquot of virusinhibitor mixture (V+I) was overlaid on MDCK cells in 96-well plates at 34°C for 1 h (Viral adsorption step). After washing with serum-free EMEM, virus-infected cells were cultured in serum-free EMEM containing 2.5 μg/ml acetylated trypsin with or without the inhibitor (at the same concentration as that in preincubation) for 18-19 h at 34°C (Treatment step). Then the cell monolayer was washed with PBS, fixed with methanol for 1 min, and incubated with anti-nucleoprotein mouse IgG (4E6) for 45 min at room temperature. β-galactosidase-labeled antimouse IgG with 0.1% block-ace was added and incubated for 45 min at room temperature. Then the enzyme reaction was started by adding 40 μM 4-methylumbelliferyl-β-D-galactoside (MU-Gal) with 50 μM MgCl 2 . After 45 min of incubation at 37°C, the reaction was stopped by addition of 100 mM sodium carbonate buffer (pH 10.6). Fluorescent intensity of released 4-methylumbelliferone (MU) was measured by excitation at 336 nm and emission at 490 nm (Ex355/Em460) using a fluorescence microplate reader (Fluorescence staining step).

Hemagglutination and hemagglutination inhibition assays
For virus quantitation, a 5-μl aliquot of virus sample serially diluted in PBS was incubated with 50 μl of 0.5% suspension of guinea pig erythrocytes for 2 h at 4°C. Virus titers expressed as HAU were determined by visual reading.
For determination of inhibition of HA binding, a 25-μl aliquot of fetuin or PVP-I serially diluted in PBS was preincubated with 25 μl of virus (4 HAU) in 96-well U-bottom plates at 4°C for 1 h. Hemagglutination was determined as described above.
To visualize agglutination directly, 100 mg/ml of PVP-I was added to a well of a 24-well flat-bottom plate to give a final concentration of 1.25, 2.50, 3.75 or 5.00 mg/ml, followed by addition of 5 μl of virus (2 14 HAU) to obtain a final HAU of about 400 [42]. To avoid possible cleavage of virus-erythrocyte binding by sialidase, erythrocyte agglutination was observed under a light microscope immediately after addition of 200 μl of 0.25% erythrocyte suspension.

Receptor binding inhibition assay
Specific binding of virus to α2,3 and α2,6 sialoglycopolymers was performed in 96-well plates coated with 0-2,000 ng/ml of sialoglycopolymers as described previously [43]. Briefly, after blocking the plates with 1% block-ace in PBS, viruses (64 HAU) in PBS containing 0.1% Tween 20 (PBST) were added and incubated for 2 h at 4°C. Viruses bound to sialoglycopolymers were detected by anti-HA mouse IgG and HRP-conjugated goat anti-mouse IgG+M, each step being conducted at 4°C for 2 h and followed by washing with PBST 5 times. The reaction color was developed by adding o-phenylenediamine (OPD) and H 2 O 2 in 100 mM citrate buffer (pH 6.0), prepared according to the instruction manual (Wako, Osaka, Japan), terminated by adding H 2 SO 4 , and assessed by measuring optical densities at 492 and 620 nm.
A receptor inhibition experiment was performed as described above, but each virus (12 HAU) was preincubated with PVP-I (0-780 μg/ml) or fetuin (0-1,250 μg/ ml) in PBST at 4°C for 2 h before the virus mixture was added to the plate coated with 50 ng/ml of sialoglycopolymer.

Sialidase inhibition assay
Five μl of 2 viral enzyme units (one unit being the amount required to liberate 1 nmole of product per min) was preincubated with 5 μl of an inhibitor serially diluted in 20 mM sodium acetate buffer (pH 6.0) at 37°C for 15 min. To determine the inhibition mechanism, 5 μl of 2 viral enzyme units was incubated with 5 μl of 280-1000 μM MUNA either alone or in the presence of 5 μl of OC, DANA or PVP-I for 15 min at 37°C. The MU products were measured as described above. Data were fitted to the Michaelis-Menten equation for determination of apparent Vmax and K m values and then transformed and plotted as Lineweaver-Burk plots using GraphPad Prism software. Inhibition constant of each inhibitor was calculated according to their inhibition type as follows: For competitive inhibitor, K mapperance = K m (1+I/K i ).
K i and K is are dissociation constants for free enzymeinhibitor and substrate-bound enzyme-inhibitor complex, respectively.


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Phenotypes induced by NM causing α-skeletal muscle actin mutants in fibroblasts, Sol 8 myoblasts and myotubes

Background Nemaline myopathy is a neuromuscular disorder characterized by the presence of nemaline bodies in patient muscles. 20% of the cases are associated with α-skeletal muscle actin mutations. We previously showed that actin mutations can cause four different biochemical phenotypes and that expression of NM associated actin mutants in fibroblasts, myoblasts and myotubes induces a range of cellular defects. Findings We conducted the same biochemical experiments for twelve new actin mutants associated with nemaline myopathy. We observed folding and polymerization defects. Immunostainings of these and eight other mutants in transfected cells revealed typical cellular defects such as nemaline rods or aggregates, decreased incorporation in F-actin structures, membrane blebbing, the formation of thickened actin fibres and cell membrane blebbing in myotubes. Conclusion Our results confirm that NM associated α-actin mutations induce a range of defects at the biochemical level as well as in cultured fibroblasts and muscle cells.

Background
Nemaline myopathy (NM) is a neuromuscular disorder, characterized by muscle weakness and hypotonia. It is a disease of the skeletal muscle sarcomere and is caused by mutations in components of the thin filament [1]; in about 20% of the cases this is in α-skeletal muscle actin (ACTA1, further referred to as α-actin) [2][3][4]. At the cellular level typical patient phenotypes are the presence of actin containing intranuclear or sarcoplasmic rod-shaped structures (nemaline rods) that may be present either in the sarcoplasm or in the nuclei [4]. Another congenital myopathy associated with ACTA1 mutations is actin myopathy, this disease is phenotypically similar to NM, and often studied together with NM, however the cellular inclusions here are actin aggregates instead of rods [4]. Curiously, the proportion of muscle fibers containing rods or aggregates does not correlate with the degree of muscle weakness [5].
To date, more than 100 α-actin mutations leading to NM or actin myopathy have been identified [6]. Given that αactin is an essential protein for muscle function it is not surprising that mutations in this protein cause diseases (reviewed in Tondeleire et al., in press, [7]). Actin molecules require folding by the chaperones prefoldin and the cytoplasmic chaperonin CCT, to become functionally active [8][9][10] and have an absolute requirement for ATP to remain stable [11]. Their self assembly results in the formation of actin filaments that in muscle are part of the thin filaments and that interact with the various actin binding proteins such as α-actinin, tropomyosin and myosin in muscle. As a result, a disease-causing mutation in actin can affect one or more of these properties or functions and this heterogeneity at the biochemical level can cause mixed phenotypes and render diseases also heterogeneous at the cellular level. This was reflected in our previous reports on several congenital myopathy causing mutants [12][13][14] and Rommelaere et al, submitted. We discovered four different biochemical phenotypes [12] and were able to induce typical disease characteristics as rods and aggregates in fibroblasts, myoblasts and myotubes [12,14]. Additionally, we found that NM associated actin mutants induce cell membrane blebbing in differentiating myoblasts (Rommelaere et al., submitted). Here we biochemically analyze 12 new α-actin mutants associated with NM and investigate their cellular phenotypes as well as further characterization of 8 mutants biochemically characterized in Costa et al. [12].

Construction and biochemical analysis of the α-actin CCD and CFTD causing mutants
Construction and biochemical analysis of the NM causing α-actin mutants were performed as described previously [12,14,15]. Briefly, mutants were expressed as 35 S-labeled proteins in in vitro transcription translation reactions in reticulocyte lysates and analysed on native gels with or without ATP, followed by autoradiography. For bandshift assays thymosin β4, DNAseI and vitamin D binding protein were added to the reaction. Co-polymerisation assays were performed by adding WT rabbit skeletal muscle α-actin. After polymerisation, the soluble and insoluble fraction were separated by centrifugation, and the amount of mutant actin in each fraction was analysed by autoradiography.

Cell culture and transfection
Cell culture and transfection were performed as previously described [12,14]. Briefly, NIH3T3 fibroblasts were transfected with lipofectamin 2000 (Invitrogen), according to manufacturer's protocol. Coverslips were mounted for immunofluorescence after 24 hours of transfection. Sol 8 myogenic cells [16] were seeded in 12 well plates containing thermanox coverslips (Nunc). After 24 hours the cells were transfected using jetPEI (Qbiogene) or nucleofected, using cell line nucleofector kit V (Amaxa biosystems) with the pcDNA3.1 vectors encoding N-terminally myc-tagged α-skeletal muscle actin (wild type or mutant). Coverslips were mounted for immunofluorescence after 48 hours of transfection. To promote fusion (F) of myoblasts into myotubes, the growth medium was replaced by a differentiation medium with 2% horse serum. Myotubes were immunostained on day F+4 or F+5.

Immunological staining
Standard immunological staining was performed [12,14]. To visualise the mutant actins we used a polyclonal antimyc antibody (Abcam) and fluorescent phalloidin (Molecular Probes) for filamentous actin. The immunolabelled samples were examined using an Olympus IX71 epifluorescence microscope and images were acquired using a cooled Spot Camera (Diagnostic Instruments) and Analysis software (Soft Imaging Systems).
For statistics of stress fibre incorporation and diffuse cytoplasmic myc-actin staining, pictures were visually inspected and blindly scored by two persons. P-values were calculated by a Fisher's exact test http:// www.socr.ucla.edu/htmls/ana/ FishersExactTest_Analysis.html on the separate experiments.

Biochemical characterization of α-skeletal muscle actin mutants causing NM
In a previous study [12], we biochemically characterized a set of actin mutants causing nemaline myopathy and discovered four different biochemical phenotypes: folding defective mutants, unstable mutants, mutants with reduced copolymerisation capacity and mutants without defects in these tests (for details and limitations of these assays see [12,15,17]). In the present study, we investigated twelve new NM mutants using similar approaches. In patient muscles, these mutants induce different abnormal structures such as intranuclear and cytoplasmic rods and/or an excess of thin filaments ( Table 1).
The locations of the mutations in the actin molecule are displayed in pink on the 3D-representation of the actin molecule ( Figure 1A) [18]. Mutations are found in sub-domains I (D25N and R116H), II (V35L, P38L, H73L and I75L), III (V163M) and IV (Q246R, G251R, R256H and M269R). Some of these residues (D25, H73, I75, R116 and V163) are in or near the nucleotide-binding pocket. [19][20][21] All these actin mutants are capable of folding properly, since they migrate at a position similar to monomeric actin in gels with ATP ( Figure 1B) and since they can bind to the ABPs tested (i.e. thymosin β4, DNAseI and VDBP, Table 1). However, for three of the mutants act E83K, R116H and V163M a significant proportion of the proteins remains bound to CCT on native gels ( Figure 1B), indicating that they fold less well compared to WT-αactin. On gels without ATP, mutants act E83K, V163M and M269R show a slight increase in CAP-binding [22]. This effect is, however, not very strong and precludes to conclusively decide that these mutants are unstable due to impaired nucleotide binding [10,17]. We note however that mutants E83K and V163M induced more aggregation in the in vitro translation reaction than does mutant R116H (Table 1). In copolymerisation tests mutants act P38L, H73L, E83K, R116H, V163M, R256H and M269R displayed reduced copolymerization capacity, whereas mutants act D25N, V35L, I75L, Q246R and G251D show normal copolymerisation.

Cellular phenotypes induced by the NEM causing αskeletal muscle actin mutants
Since we previously observed that expression of myctagged α-actin mutants associated with NM can induce a range of defects in the actin cytoskeleton, we constructed all of these mutants with an N-terminal myc-tag and transfected them in NIH3T3 fibroblasts and Sol8 myob-lasts [12,14] and Rommelaere et al. submitted. In order to analyse their incorporation in the actin cytoskeleton or the occurrence of rods or aggregates, we performed immunological staining of the myc-tagged mutants, together with fluorescent phalloidin staining for F-actin containing structures. We performed a similar analysis on myc-tagged actin mutants N115S, G182D, R183G, R183C, Q263L and G268C (green in Figure 1A), which were previously biochemically characterized in Costa et al. (Table 2) (additionally, G182D which was only characterized in fibroblasts in Costa et al. was analysed here in muscle cells) [12]. The results of this ex vivo analysis are summarized in Table 3.
Upon expression of act V163M in myoblasts we also observed intranuclear rods. Another mutation at this position, V163L, similarly induces intranuclear rods in both fibroblasts and myoblasts [12]. V163 lies close to the nuclear export signal [23] and therefore this mutation could interfere with nuclear export resulting in intranuclear rods (Figure 2). Domazetovska et al. expressed EGFPor untagged V163L and V163M actin in fibroblast and myoblast cell lines and observed similar phalloidin positive rods. Our co-polymerization results support their observation that intranuclear rods induced by V163M The properties investigated here are folding, binding to cyclase associated protein (CAP), binding to the actin binding proteins (ABP) thymosinβ4, DNAseI and Vitamin D binding protein (VDBP) and copolymerisation with rabbit α-skeletal muscle actin (copol). Also the location in the actin structure according to Kabsch et al. [18] is indicated, in addition to the patients phenotype as presented in the reference with NEM = nemaline myopathy, sarcoplasmic nemaline bodies; IRM = intranuclear rod myopathy.
(A) Location of the studied ACTA1 mutations on the 3D-representation of the actin molecule [18] Figure 1 (A) Location of the studied ACTA1 mutations on the 3D-representation of the actin molecule [18]. Mutations indicated in green were biochemically analysed in this study, mutations indicated in pink were analysed in Costa et al. [12]. (B) Native gel analysis of twelve α-actin mutants causing NM. Autoradiograms of native gels of 35 S-labeled WT α-actin and 12 mutants causing NM run with or without ATP. Actins were produced in in vitro transcription translation reactions in reticulocyte lysate, which endogenously contains CAP [22] and the actin folding machinery, prefoldin (PFD) and CCT [8,9,15]. The positions of these complexes with actin are indicated. The graph underneath indicates the percentage of actin in each of the bands (associated with CCT (purple), with prefoldin (white, (+ATP)) or CAP (white, (-ATP)) and free actin (red). In each lane the total actin is the sum of these three actin species.  might be due to an increased availability of monomeric actin caused by reduced incorporation of the mutant in actin filaments [24,25]. However we can not generalize this to the other mutants as I75L for instance displays normal co-polymerisation.
Another phenotype associated with expression of nemaline myopathy associated actin mutants in cultured cells is the formation of actin-containing phalloidin negative aggregates. For eleven of the investigated mutants we observed these in fibroblasts (D25N, E83K, R116H, V163M, R183G, R183C, Q246R, G251D, G268C and M269R) (Figure 3). In most cases the aggregates were cytoplasmic or perinuclear, yet for R116H and V163M we additionally observed intranuclear aggregates, consistent with the fact that these mutants also induce intranuclear rods. Strikingly, E83K, R116H and V163M also aggregate in the in vitro translation assays (see above). In addition, for a subset of mutants, we also observed cytoplasmic aggregates in Sol8 myoblasts. This is the case for mutants D25N, P38L, I75L, M132V, V163M, R183C, G251D, R256H and Q263L ( Figure 3).
As in previous reports [12], not all mutants induced the formation of rods or aggregates, and these phenotypes are induced more in fibroblasts than in muscle cells. This supports the hypothesis that rods can be considered secondary effects in patient muscles, which serve as 'rescue' mechanisms to remove the mutant actin from the active pool, and that other defects could underlie the muscle weakness in NM patients [26] (Rommelaere submitted).
The fact that we observed both phalloidin positive and negative rod-like structures and aggregates, is consistent with previous reports on fibroblasts and muscle cells expressing NM causing actin mutants [13,27]. A lack of phalloidin staining could indicate that the rods do not contain filamentous actin, or that the F-actin is not accessible for the phalloidin. At present it is not known whether patient rods stain with phalloidin however our results indicate that rods induced by different mutants could have a different composition and/or organisation.

Diffuse cytoplasmic myc-staining in fibroblasts
To test the ability of the mutants to incorporate in filamentous structures, we calculated the percentage of fibroblasts in which a particular mutant is able to incorporate in stress fibres. In addition, we also scored for diffuse cytoplasmic staining of the myc-tagged mutant, which gives a second indication of the (lack of) incorporation in filamentous structures. Cells were visually inspected and blindly scored for both parameters by two persons and the mean was calculated. Similar to WT, all mutants are able to incorporate in stress fibres in a certain percentage of the cells. For G182D and I375L actin expressing fibroblasts this was significantly lower than for WT actin expressing cells ( Figure 4A). The percentage of cells with a diffuse cytoplasmic myc-staining, shows more variation ( Figure  4B): V35L, P38L, H73L, and R183G induce this cytoplasmic staining in significantly more cells than WT mycactin. N115S, V163M, G182D, Q246R, G251D, R256H, M269R and I375L also have a tendency to display this phenotype more than cells expressing WT myc-actin, yet they do so to a lesser extent than the former mutants.

Five mutants induce cell blebbing in muscle cells
Consistent with our previous observation that particular NM actin mutants induce blebbing in myoblasts (Rommelaere et al., submitted), we also found five mutants here displaying this phenotype: act P38L, E83K, R116H, R183C and Q246R ( Figure 4C-F). Each of these mutants is enriched in the blebs.

Phenotypes induced in myotubes
We differentiated myoblasts, transfected with the α-actin mutants, during five days and allowed them to form multinucleated myotubes. Immunological staining revealed that all studied mutants incorporate to a certain degree in actin fibers in myotubes (shown for H73L and G268C in Figure 5). Act R116H, M132V, V163M transfected myotubes show an aberrant phenotype at this differentiation stage ( Figure  5). In myotubes expressing act V163M, actin fibers appear thickened, a phenotype we, and others, previously also observed for act V163L (Rommelaere et al., submitted, [13]). This cellular phenotype is also consistent with the sarcomeric disorganization in Drosophila muscles expressing V163M [25]. Act R116H and M132V expressing myotubes display blebs or membranous protrusions as described for other NM mutants by Rommelaere et al. (Figure 5).

Conclusion
We corroborate that NM associated α-actin mutations can induce a range of defects, both at the biochemical and cellular level. We show that the twenty NM mutants investigated here score at least one of the previously described cellular NM or actin myopathy associated phenotypes (Table 3), i.e. the cells display intranuclear or cytoplasmic rods, aggregates, blebs (only in differentiating myoblasts), diffuse cytoplasmic myc-actin staining or thickened actin fibres in myotubes. As observed before, not a single mutant can reproduce all phenotypes although some mutants display multiple, occasionally cell line dependent, phenotypes. On the other hand, not all mutants induce rod-like structures or aggregates, hallmarks in patient muscles, which supports the fact that rods could be secondary effects, and that other mutant induced defects are underlying the muscle weakness in NM patients. Phalloidin staining of cells also indicate that nemaline rods induced by different actin mutants can have different organisation or composition. Additionally, as concluded before in Costa et al., [12] there is no correlation between the observed biochemical and cellular phenotypes nor between one of these and the clinical situation.
α-actin mutants induce rods in cell line cultures