Genetics of human hydrocephalus

Human hydrocephalus is a common medical condition that is characterized by abnormalities in the flow or resorption of cerebrospinal fluid (CSF), resulting in ventricular dilatation. Human hydrocephalus can be classified into two clinical forms, congenital and acquired. Hydrocephalus is one of the complex and multifactorial neurological disorders.
Introduction
Human hydrocephalus is a significant medical condition with an estimated incidence of 1 in 1500 births [1]. Hydrocephalus is characterized by abnormalities in the flow or resorption of cerebrospinal fluid (CSF), resulting in ventricular dilatation. However, hydrocephalus is far more complicated than a simple disorder of CSF circulation [2]. Although commonly considered a single disorder, human hydrocephalus is a collection of a heterogeneous complex and multifactorial disorders [3]. Genetic factors are involved in the pathogenesis of hydrocephalus [4–6]. For the purposes of this review, we categorize hydrocephalus as congenital, which is present at birth and often associated with developmental defects; and acquired, which occurs after development of the brain and ventricles [7–12].
The development and progression of congenital hydrocephalus is a dynamic process that is not yet well understood. It is thought that it may develop at an important and specific embryonic time period of neural stem cell proliferation and differentiation in the brain [13, 14]. Congenital hydrocephalus may occur alone (non-syndromic) or as part of a syndrome with other anomalies (syndromic) [15, 16]. In syndromic forms, it is hard to define the defective gene because of the association with other anomalies. We will mainly focus on isolated forms of hydrocephalus. In genetic terms, the isolated (non-syndromic) form of hydrocephalus is a primary and major phenotype caused by a specific faulty gene.
It is estimated that about 40% of hydrocephalus cases have a possible genetic etiology [10]. In humans, X-linked hydrocephalus (HSAS1, OMIM) comprises approximately 5–15% of the congenital cases with a genetic cause [10, 17–20]. Although there is strong evidence for genetic causes, only one hydrocephalus gene (X-linked) has been identified in humans.
Besides genetic factors, many other factors influence the development of congenital hydrocephalus, such as congenital malformations, intracerebral hemorrhage, maternal alcohol use [21, 22], infection [6, 23–25], and X-ray radiation during pregnancy [26, 27].
Genetics in hydrocephalus
Congenital hydrocephalus is the more common of the two forms of hydrocephalus, and is probably the consequence of abnormal brain development and perturbed cellular function, which emphasizes the important roles that congenital hydrocephalus genes play during brain development. In general, the recurrence risk for congenital hydrocephalus excluding X-linked hydrocephalus is low. Empiric risk rates range from <1% to 4% [28–30], indicating the rarity of autosomal recessive congenital hydrocephalus [10, 20, 31, 32]. However, multiple human kindreds with congenital hydrocephalus have been reported [10, 15, 20, 32–43]. The loci or genes for human autosomal recessive congenital hydrocephalus have not yet been identified, but there is at least one locus for this trait. Furthermore, like animal models, since there is heterogeneity among clinical phenotypes, there may be more genetic loci in human autosomal recessive congenital hydrocephalus.
One kindred was reported in which congenital hydrocephalus was transmitted in an autosomal dominant fashion. This condition was associated with aqueductal stenosis but was not associated with mental retardation or pyramidal tract dysfunction. The lack of mental retardation and pyramidal tract dysfunction was in contrast to X-linked or recessive congenital hydrocephalus with stenosis of the aqueduct of Sylvius (HSAS), in which these abnormalities are commonly seen [44]. Another study identified a kindred with a microdeletion of 8q12.2-q21.2 which subsequently developed hydrocephalus. This trait was also transmitted in an autosomal dominant fashion [45]. Molecular genetic studies have revealed that the responsible gene for X-linked human congenital hydrocephalus is at Xq28 encoding for L1CAM (L1 protein) [46]. The mutations are distributed over the functional protein domains. The exact mechanisms by which these mutations cause a loss of L1 protein function are still under investigation.
Another form of this disorder, acquired or adult-onset hydrocephalus is mostly sporadic and characterized by ventricular enlargement in the absence of significant elevations of intracranial pressure; therefore this form is termed normal pressure hydrocephalus (NPH). Definite changes in CSF flow, resorption, and associated dynamics have been found in NPH patients, and these changes may represent a pathogenic mechanism or a secondary phenomenon [47]. Adult-onset hydrocephalus may develop either as a result of decompensation of a “compensated” congenital hydrocephalus, or it may arise de novo in adult life secondary to an acquired disturbance of normal CSF dynamics. The latter may be due to late-onset aqueductal stenosis or disruption of normal CSF absorptive pathways [11, 48]. Acquired (adult-onset, or NPH) form of inherited hydrocephalus is very rare. Recently, an X-linked adult-onset NPH [49] and a form of familial NPH that is transmitted in autosomal dominant fashion [50] have been reported, but detailed genetic linkage studies have not been carried out yet. The genetic etiology of this form is therefore totally unknown.
Hydrocephalus has been observed in many mammals [51–59]. Animal hydrocephalus models have many histopathological similarities to humans and can be used to understand the genetics and pathogenesis of brain damage [59–64]. It has been well documented in the animal models that in the majority of cases, congenital hydrocephalus is a genetic disease. Furthermore, many congenital hydrocephalus loci have been mapped and identified in the animal models.
Hydrocephalic Texas strain (HTX) rat model of inherited congenital hydrocephalus is characterized by onset in late gestation, a complex mode of inheritance, and ventricular dilatation associated with abnormalities in the cerebral aqueduct and subcommissural organ (SCO), a structure that is important for the patency of the aqueduct of Sylvius and normal CSF flow in the brain. Quantitative trait locus (QTL) genetic mapping has been performed from the progeny of a backcross of HTX rat with the non-hydrocephalic Fischer F344 strain. The disease has been linked with loci on chromosome (Chr) 9 (peak markers D9Rat2), 10 (between markers D10Rat136 and D10Rat135), 11 (peak markers D11Arb2 and D11Rat46) and 17 (peak markers D17mit4 and D17Rat154) respectively. The severity of hydrocephalus in HTX rat seems to be influenced by different genetic loci [65–68]. Another study suggested that the HTX strain is homozygous carrier of an autosomal recessive hydrocephalus gene with incomplete penetrance [69]. The genetics of another hydrocephalus inbred strain, Wistar-Lewis rats (LEW/Jms) which demonstrate inherited congenital hydrocephalus, is less clear with possible traits as an autosomal recessive [70] or semidominant or multigenic (possible QTL) with a possible locus on sex chromosomes [71], but none of the loci has been localized.
In mouse models, three QTL loci associated with congenital hydrocephalus have been identified and labeled as Vent8a, Vent4b, and Vent7c. As a major QTL controlling variance in ventricular size, Vent8a is located on Chr 8 (near the markers D8Mit94 and D8Mit189). The Other two loci, Vent4b and Vent7c, show strong epistatic interactions affecting ventricular size in the developing embryo. Vent4b is located on Chr 4 (near D4Mit237 and D4Mit214), and Vent7c is located on Chr 7 (between D7Mit178 and D7Mit191) [72].
The autosomal recessive congenital hydrocephalus-1 (hy1) mouse has been characterized phenotypically by a dome-shaped head that is sometimes seen at birth or develops during the first 2 weeks. Internally, dilatation of the entire ventricular system is observed [73, 74]. A more severe phenotypic form, hydrocephalus-2 (hy2) mouse [75, 76], and an obstructive hydrocephalus (oh) mouse with communicating hydrocephalus and secondary aqueductal stenosis have also been described [77, 78]. Unfortunately subsequent efforts to identify genetic loci have not been done on these non-inbred mouse strains.
In mouse targeted insertional mutagenesis, the accidental insertion of a transgene into a crucial genomic locus could yield important information, which has happened twice in hydrocephalus genetic studies. The transgenic mouse line OVE459 demonstrates autosomal recessive congenital hydrocephalus. This is caused by a Bdnf transgene-induced insertional mutation on a single locus on mouse Chromosome 8 (near marker D8Mit152). The OVE459 insertion locus is overlapped with that of autosomal recessive hydrocephalus-3 (hy3) mouse that phenotypically shows lethal communicating hydrocephalus with perinatal onset [79, 80]. The transgene insertion resulted as a rearrangement of Hydin exons in OVE459 mice. Subsequently, a single CG base-pair deletion in exon 15 of Hydin was also discovered in hy3 mice carrying the spontaneous hy3 mutant allele [81, 82].
In another targeted insertional mutagenesis resulting in congenital hydrocephalus, the CYP2J2 transgene interferes with the expression of a brain-specific isoform of the regulatory factor X4 (RFX4), which belongs to the winged helix transcription factor family. This brain specific isoform is called variant transcript 3 or RFX4_v3 and is crucial for normal brain development as well as for the genesis of the SCO. Loss of a single allele prevents formation of the SCO and leads to an autosomal dominant congenital hydrocephalus. This obstructive hydrocephalus appeared to be secondary to failure of development of the SCO [83].
The autosomal recessive congenital hydrocephalus (ch) mouse was reported decades ago [79]. Recently this mouse has been shown to have a mutation on another winged helix/forkhead transcription factor gene, Foxc1 (Mf1) on mouse Chromosome 13 [84, 85]. There is a recent report of 6 children with hydrocephalus from 3 different families with subtelomeric deletions from chromosome 6p. Three forkhead genes within this region (FOXF1 and FOXQ1) or proximal to it (FOXC1) were evaluated as potential candidate disease genes but no disease causing mutations were identified [86].
The autosomal recessive hydrocephalus with hop gait (hyh) mouse exhibits dramatic dilation of the ventricles at birth and invariably develops hopping gait. The hyh mouse shows a markedly small cerebral cortex at birth and dies postnatally from progressive enlargement of the ventricular system. The small cortex in hyh mouse reflects altered development of the neuronal cells. In this mouse, it is postulated that neural progenitor cells withdraw prematurely from the cell cycle, producing more early-born, deep-layer cerebral cortical neurons but depleting the cortical progenitor pool, and creating a small cortex. Genetic linkage analysis localized the hyh locus between markers D7Mit75 and D7Mit56 on mouse Chr 7. Later, the hyh gene was identified as α-SNAP (soluble NSF attachment protein α) [87]. Homozygous mutant mice harbor a missense mutation M105I in a conserved residue in one of the α-helical domains. The hyh mutant was not a null allele and is expressed; however, the mutant protein is 40% less abundant in hyh mice.
The autosomal recessive hemorrhagic hydrocephalus (hhy) homozygous mutant mouse has dilated lateral ventricles and a patent aqueduct, with no histological abnormalities either in the subarachnoid space or in the choroid plexus. Multiple hemorrhages in the meninges and throughout the brain parenchyma can be observed in the advanced stages of hydrocephalus. The hhy locus has been localized on mouse Chr 12 [88].
Recently, several new congenital hydrocephalus models have emerged in zebra fish mutagenesis screening. These models have been shown to have the defects in embryogenesis and early development leading to enlarged brain ventricles. However, genetic loci for these models have not been identified yet [89, 90].
Genetic studies in animal models have started to open the way for understanding the underlying pathology of hydrocephalus. In contrast to research with animal models, human hydrocephalus genetic research has lagged far behind. To date, at least 43 mutants of hydrocephalus have been described, and 10 congenital hydrocephalus genes have been identified. Among them, only one hydrocephalus gene has been identified in humans (see Table 1).
Table 1Summary of current known loci (or mutants) of hydrocephalus in vertebratesSpeciesStrainClinical formTrait*LocusChromosomeHuman syntenic regionHuman GeneReferencesHumanCARUnknownunknown10,15,27,32–43HumanCADUnknown8q12.2–21.2 or unknown 44,45HumanAOADNPHunknown50HumanCX-linkedL1camXXL1CAM46HumanAOX-linkedUnknownXX49RatHTXCQTLD9Rat2 9q38 5q21.1, 18p11.22–31 65–68RatHTXCQTLD10Rat136, D10Rat13510q32.1–10q32.317q21.3–q25.365–68RatHTXCQTLD11Arb2, D11Rat4611q233q27–28, 22q11.21,10p12.2 65–68RatHTXCQTLD17Mit4, D17Rat15417q12.1 1q43, 10p11.21–p13 65–68RatLEW/JmsCAR, (QTL)unknownunknown70,71MouseC57BL/6JCQTLVent8a88p11–23, 13q11–3472MouseC57BL/6JCQTLVent4b46p, 972MouseC57BL/6JCQTLVent7c719q10–1372Mousehy1CARunknownunknown73,74Mousehy2CARunknownunknown75,76Mousehy3CARHydin816q22.2HYDIN79–82MouseC57BL/10JCARhyh719q13.3a-SNAP87, 100MouseC57BL6/JCADRfx41012q24RFX483MouseBALB/cHeACARhhy1214q3288MousechCARMf1136p25FREAC-384MouseSTOCK tbCARohunknown77,78MouseC57BL/6*CBA/JCARMdnah5155p15.2DNAH5106MouseC57BL/CBACADOtx21414q21-q22OTX2122Mouse129P2/OlaHsdCARMsx154p16.3-p16.1MSX1107,108MouseC57BL/6CARSocs71117q12SOCS7118MouseC57BL/6JCARNmhc-b1117q13MYH10121Zebrafishm404/m491CARapounknown89,90Zebrafishm409/m432CARcudakunknown89,90Zebrafishm691CAReagleunknown89,90Zebrafishm591CARendeavorunknown89,90Zebrafishm584CARenterpriseunknown89,90Zebrafishm492/m510CARgalileounknown89,90Zebrafishm445/m585/m700CARgumowyunknown89,90Zebrafishm727CARhubbleunknown89,90Zebrafishm221/m470/m680CARinterrailunknown89,90Zebrafishm733CARkeplerunknown89,90Zebrafishm728CARneilunknown89,90Zebrafishm481CARpan twardowskiunknown89,90Zebrafishm172/m476CARuchu hikoushiunknown89,90Zebrafishm766CARvoyagerunknown89,90Zebrafishm331CARvikingunknown89,90Zebrafishm479/m627CARyuraunknown89,90Zebrafishm111/m307/m512/m97CARzezemunknown89,90* Genetic trait, AR: autosomal recessive, AD:autosomal dominant, QTL: Quantitative trait locus, ** clinicalform, C: congenital, AO: Adut-Onset
Developmental, physiological and anatomical pathology of hydrocephalus
The neuropathology of hydrocephalus has been adequately elucidated. Cerebral ventricle dilatation secondary to disturbed CSF flow has been observed as an inheritable trait in a variety of laboratory animals (as well as in humans). In most cases, defective development of the cerebral aqueduct or  the subarachnoid space has been observed [61]. Affected individuals may have severe developmental delay and radiographic findings of hydrocephalus [91].
The morphological and developmental changes in the ventricular system have been well studied in three major rat models of congenital hydrocephalus: 6-aminonicotinamide (6-AN)-induced, LEW/Jms and HTX mutant rats. Comparative morphological studies revealed that 6-AN-induced hydrocephalus was comparable to the Dandy-Walker syndrome. The LEW/Jms and HTX mutant models were identical with regard to the form of presentation and progression of hydrocephalus in the postnatal period; but the pathogenesis of these two conditions in the fetal period was different. The LEW/Jms rats showed primary congenital aqueductal stenosis in early prenatal life and the hydrocephalic state appeared before pulmonary maturation was completed. However, although the model has been considered to be of congenital communicating hydrocephalus [64], the HTX fetuses demonstrated secondary closure of the aqueduct in the perinatal period. This secondary closure of the aqueduct in HTX rats is believed to be due to retrograde degeneration of the thalamus caused by apoptotic cell death [92, 93] and failure in cell proliferation [94, 95]. The HTX rat also shows a reduction in the secretory cells of the SCO. Regarding the role of the SCO in hydrocephalus pathogenesis, serial brain sections through aqueduct regions containing the SCO from HTX rats, in comparison with normal Fischer F344 strain, have been studied and found that reduced SCO glycoprotein immunoreactivity precedes both aqueduct closure and expansion of the lateral ventricles in the HTX rate (as it’s redundant) [96, 97].
Although some studies have addressed the activation of macrophages and microglia (the resident mononuclear phagocytes of the brain) within the brain in animal hydrocephalus models, little is known of their state of activation or regional distribution in human congenital hydrocephalus. In one experiment, brain tissue samples of 10 human fetal cases with hydrocephalus and 10 non-hydrocephalic controls were stained immunohistochemically with antibodies directed against MHC class II and CD68 antigens, and lectin histochemistry was done with tomato lectin. Hydrocephalus cases showed focal collections of CD68 and tomato lectin-positive macrophages along the ependymal lining of the lateral ventricles, particularly within the occipital horn. By comparison, brain tissue samples from controls showed few or no ependymal or supraependymal macrophages and the few macrophages that were present were not as intensely immunoreactive as in the hydrocephalus cases. The macrophage response detected at the ependymal lining of the ventricles and within the periventricular area in hydrocephalus may be related both to the severity of hydrocephalus and the age of the fetus [98]. Microglia that are normally interspersed throughout the intermediate zone and circumscribing the basal ganglia were within normal confines in all cases examined. Unexpectedly, hydrocephalic cases also showed focal regions of hypovascularization or alterations in the structure and orientation of capillaries within periventricular areas, compared with controls [98].
In summary, the pathological studies of hydrocephalus clearly indicate that impaired and abnormal brain development in the early development stage caused by altered neural cell fate and perturbed regulation of cellular proliferation and apoptosis. The abnormal brain development subsequently leads to the accumulation of the CSF in cerebral cavities. All these cellular and developmental events eventually lead to the congenital hydrocephalus accompanied by possible secondary inflammatory reaction and neurovascular pathogenesis.
The molecular and cellular etiology of hydrocephalus
One of the possible mechanisms leading to the pathogenesis of hydrocephalus is the disruption of neural cell membrane proteins that play an important function during brain development. The L1 protein coded by human X-linked hydrocephalus gene is a member of the immunoglobulin superfamily of neural cell adhesion molecules that is expressed in neurons and Schwann cells, and seems to be essential for the brain development and function. Hirschsprung’s disease (HSCR) is characterized by the absence of ganglion cells and the presence of hypertrophic nerve trunks in the distal bowel. There have been several reports of patients with X-linked hydrocephalus and HSCR with a mutation in the L1CAM gene. Therefore, decreased L1CAM may also be a modifying factor in the development of HSCR [99].
The gene carrying the mutation for autosomal recessive hydrocephalus in the hyh mouse codes α-SNAP protein. α-SNAP is essential for apical protein localization and cell fate determination in neuroepithelial cells [100]. α-SNAP plays a key role in a wide variety of membrane fusion events in eukaryotic cells. Membrane fusion is required for two main cellular functions: 1) the transport of molecules to distinct inter- and intracellular compartments and thereby maintenance of the functional and structural organization of eukaryotic cells; 2) the intercellular communication such as the regulated exocytosis (secretion) of neurotransmitters by neuronal cell, which occurs temporally and spatially as the precise sequential regulation events at the plasma membrane during the early brain development [87]. In the hyh mouse model, altered neural cell fate is also accompanied by abnormal localization of many apical proteins implicated in regulation of neural cell fate, including E-cadherin, beta-catenin, atypical protein kinase C (aPKC), inactivation-no-afterpotential D-like (INADL), SNAP receptor (SNARE), and vesicle-associated membrane protein-7 (VAMP-7) [100]. Furthermore, disturbed astrocyte metabolism in the early brain development in the kaolin-induced rat model of hydrocephalus has also been reported [101].
Hydrocephalus may also be caused by a malfunction of the ependymal cells [102–105]. Within the neonatal brain of the mouse, the hy3 protein (hydin) is confined to the ciliated ependymal cell layer lining the lateral, third and fourth ventricles. Hydin is not closely related to any previously known protein, with the exception of a 314 amino acid domain with homology to caldesmon, an actin-binding protein, suggesting that hydin interacts with the cytoskeleton [81, 82]. The protein of axonemal heavy chain 5 gene (Mdnah5), dynein is also specifically expressed in ependymal cells, and is essential for ultra structural and functional integrity of ependymal cilia. In Mdnah5-mutant mice, lack of ependymal flow causes closure of the aqueduct and subsequent formation of triventricular hydrocephalus during early postnatal brain development. The higher incidence of aqueductal stenosis and hydrocephalus formation in patients with ciliary defects proves the relevance of this novel mechanism in humans [106].
Hydrocephalus may be caused by malfunction of mesenchymal cells. In mice, Msx1 is a regulatory gene involved in epithelio-mesenchymal interactions in limb formation and organogenesis. In the embryonic brain, the Msx1 gene is expressed along the dorsal midline. The most important features observed in homozygous Msx1 mutants were the absence or malformation of the posterior commissure (PC) and of the SCO, the collapse of the cerebral aqueduct, and the development of hydrocephalus. The heterozygous mutants developed an abnormal PC and smaller SCO, as revealed by specific antibodies against SCO secretory glycoproteins. About one third of the heterozygous mutants also developed hydrocephalus; therefore the phenotype may be determined by the Msx1 gene dosage during a specific developmental period [107, 108]. In the autosomal recessive congenital hydrocephalus (ch) mouse model, a truncated protein lacking the DNA-binding domain of the forkhead/winged helix gene, Mf1, was generated. Mesenchymal cells from Mf1lacZ embryos differentiate poorly into cartilage in micromass culture and do not respond to added bone morphogenetic protein 2 (BMP2) and transforming growth factor-beta 1 (TGFB1). The differentiation of arachnoid cells in meninges of the mutant mice is also abnormal. The levels of developmental growth factors such as TGFB1 and BMP2 are dramatically increased in the ch mouse, and it is possible that phenotypic hydrocephalus of Mf1 mouse is due to the secondary effect of these elevated growth factors. Corresponding to studies in the ch mouse, human patients with deletions in the region containing human Mf1 homolog FREAC3 were found to develop multiple developmental disorders, including hydrocephalus [84]. Another winged helix transcription factor causing congenital hydrocephalus when mutated, RFX4_v3 transcript, is dynamically expressed in the developing brain from the neural plate stages. The RFX proteins belong to the winged-helix subfamily of helix-turn-helix transcription factors, and bind to ‘X-boxes’ in target DNA sequences and regulate expression of the downstream target genes [83]. Disruption of both RFX4_v3 alleles by insertional mutagenesis severely alters early brain morphogenesis, reduces Msx2 expression, and causes a deficiency in WNT signaling [83]. This may suggest that RFX4_v3 is probably upstream of Mf1 in the signaling pathway during early brain development.
Hydrocephalus may be caused by perturbation of growth factor signaling [109, 110]. Developmental abnormalities in congenital hydrocephalus provide the clues for the perturbation of major signaling pathways in the development [111]. TGFB is an important cytokine and growth-signaling molecule in the brain. In mouse models, severe hydrocephalus has been observed in transgenic mouse overexpression of TGFB1 in astrocytes [112, 113]. In the HTX rat, increased level of TGFB3 may contribute to the development of hydrocephalus [114]. In mouse models, fibroblast growth factor-2 (FGF-2) seems to play a predominant role in the proliferation of neuronal precursors and in neuronal differentiation in the developing cerebral cortex even at relatively late stages of brain neurogenesis. Administration of FGF-2 to embryonic brain induces hydrocephalic brain morphology and aberrant differentiation of neurons in the postnatal cerebral cortex [110]. IGF binding protein-1 (IGFBP-1) modulates the cellular action of insulin-like growth factors (IGFs), some of which are expressed in the fetal brain. Hydrocephalus has been observed in mouse models that overexpress liver-specific IGFBP-1 during fetal life. The hepatic over-expression of IGFBP-1 may have endocrine effects on brain development and induction of congenital hydrocephalus [115]. Other studies have shown that up-regulation of certain growth factors in the brain could lead to altered brain fluid dynamics [116, 117]. SOCS7 is a member of the suppressor of cytokine signaling (SOCS) protein family. SOCS proteins have a similar structure: an N-terminal domain of variable length, a central Src homology-2 domain, and a C-terminal SOCS box. Biochemical and genetic studies have revealed that SOCS family members play an important role in the termination of cytokine and growth factor signaling. Homozygous Socs7 mutant mice were born in expected numbers, were fertile, and did not exhibit defects in hematopoiesis or circulating glucose or insulin concentrations. Strikingly however, these homozygous Socs7 mice were 7–10% smaller than their wild-type littermates, and within 15 weeks of age approximately 50% of the homozygous Socs7 mice died as a result of hydrocephalus. In situ hybridization studies in normal mice have revealed that Socs7 is prominently expressed in the brain, suggesting that SOCS7 plays an important functional role in early brain development [118]. We can therefore hypothesize that loss of SOCS7 function will lead to increased expression of cytokines resulting in developmental abnormalities and congenital hydrocephalus due to its inhibitory role in cytokine signaling.
Hydrocephalus may also be caused by the disruption of extracellular matrix (ECM). In the TGFB1 over-expression mouse model, the changing expressions of a remodeling protein - matrix metalloproteinase-9 (MMP-9) and its specific inhibitor- tissue inhibitor of metalloproteinases-1 (TIMP-1) were also found to be important factors in the spontaneous development of hydrocephalus by altering the ECM environment [119]. Furthermore, increased expression of cytokines such as TGFB1 might also reciprocally play an important role by disrupting the vascular ECM remodeling, promoting hemorrhages, and altering the re-absorption of CSF [120]. In another mouse model, ablation of the nonmuscle myosin heavy chain II-B (NMHC-B) results in severe hydrocephalus with enlargement of the lateral and third ventricles. These defects may be caused by abnormalities in the cell adhesive properties of neuroepithelial cells and suggest that NMHC-B is essential for both early and late developmental processes in the mammalian brain [121].
Hydrocephalus may also be caused by the disruption of major early brain developmental patterning molecules. Autosomal dominant hydrocephalus in Otx2 mutation mice is characterized by eminent dilatation of lateral ventricles and a ballooned cerebrum. Histopathology shows edematous change of the periventricular white matter, suggesting that Otx2 functions as a brain developmental organizer, and a disruption of this gene is a likely cause of hydrocephalus [122].
In conclusion, many genetic loci of hydrocephalus have been defined in animal models, which is building a foundation for better understanding of molecular etiology of hydrocephalus; however, genetic research of hydrocephalus in humans is limited. The histopathological similarities of animal models can be used to understand the genetics and pathogenesis of human hydrocephalus. For example, the histopathological and morphological appearance of hydrocephalic HTX rats is demonstrated in Fig. 1, in which hydrophilic HTX rat exhibits large cerebral ventricles, a progressively-thinned cortical mantle, and stretched internal capsule fibers[123]. Review of the molecular etiologies shows a very diverse set of pathogenetic mechanisms. Perturbation of almost any molecule that plays a crucial role in early brain development, and sequentially regulation of dynamics of cerebrospinal fluid, could lead to the pathogenesis of congenital hydrocephalus. The 10 known hydrocephalus genes mostly code for important cytokines, growth factors, or related molecules in the cellular signal pathways during early brain development.
Fig. 1Comparison of rat brain morphology by MRI and histology at 4, 11 and 21 days of age in non-hydrocephalic and hydrocephalic HTX rats. T2-weighted MRI scans of coronal sections from a non-hydrocephalic HTX rat at the level of the thalamus at 4, 11 and 21 days (A, D, G) shows small ventricular and subarachnoid spaces compared to an age-matched HTX littermate (B, E, H) that exhibits large cerebral ventricles, a progressively-thinned cortical mantle, and stretched internal capsule fibers (*). Congenital closure of the cerebral aqueduct becomes life-threatening by 21 days of age. Histological sections (C, F, I) at the level of the midbrain at the same ages demonstrate extreme thinning of the cortical mantle. MRI images are modified from Jones et al (2000) [123] with permission by Maney Publishing; histological samples are from the doctoral thesis of Janet M. Miller, PhD
Future prospects
It is essential to recognize that molecular genetics is the only current scientific approach that can be used to study hydrocephalus in which the usual concern about whether an observed phenomenon is a consequence or a cause is completely addressed.
Despite our knowledge of the genetics of hydrocephalus in animal models, we have very limited knowledge about the genetic and molecular mechanisms that cause human hydrocephalus. Without this knowledge, it is impossible to say whether the pathogenesis of human hydrocephalus is comparable to that seen in animal models, and impractical to extrapolate data gained from animal models to humans. In order to better understand human hydrocephalus and to develop more appropriate translational research, it will be necessary to conduct large-scale genetic studies of human hydrocephalus. If, and when, more heritable forms of human hydrocephalus are identified, and underlying genes and their functions are characterized, then this knowledge could be used to improve patient care in a variety of different ways such as prenatal diagnosis and new potential therapeutic approaches. Possible new mechanisms other than altered CSF circulation and resorption, if uncovered via the genetic research, may also help explain why patients with hydrocephalus may experience symptomatic progression despite functioning shunts.
Efforts to identify genetic variants associated with susceptibility to genetic diseases rely on three major approaches: pedigree and sib-pair linkage analysis and population association studies. The differences among these study designs reflect their derivation from biological versus epidemiological traits. Like most common diseases, it would be very difficult to identify and recruit large pedigrees in hydrocephalus that show hereditary transmission of the condition. Therefore, the last two approaches, sib-pair linkage analysis and population association studies, are the best options for the genetic study on this disease. For any study, but particularly in the case of genetic mapping for a common disease, a large sample size is crucial in achieving statistical significance. Recent advances in the genomics and statistical methodology in genetic mapping will certainly help in making well-powered studies more feasible, by reducing the number of genetic markers or workload required for these studies. For example, since many genes and loci response for hydrocephalus in animal models have been mapped, candidate genes approach will certainly be the very first choice to test the collected human hydrocephalus population for linkage and association analysis.
In collaboration with the Hydrocephalus Association (HA), our group has initiated a genetic study of human hydrocephalus. As part of a prospective study that has been approved by the Johns Hopkins Institutional Review Board, we are collecting blood samples from both congenital and acquired NPH hydrocephalus patients. The objectives of this study are to identify the genetic loci responsible for the development of hydrocephalus, to examine the relationship between genotype and phenotype and to define the functions of these genes during early development. This is the first large-scale research study of its kind and information gained from this study will undoubtedly provide invaluable information concerning the developmental mechanisms of this disease in humans. Such knowledge will hopefully lead to the more reasonable treatment schemes, the better diagnostic tools, and the more effective therapeutic modalities.

genetic of
hydrocephalus
congenital
acquired
multifactorial disorder


Muscle wasting in chronic kidney disease: the role of the ubiquitin proteasome system and its clinical impact

Muscle wasting in chronic kidney disease (CKD) and other catabolic diseases (e.g. sepsis, diabetes, cancer) can occur despite adequate nutritional intake. It is now known that complications of these various disorders, including acidosis, insulin resistance, inflammation, and increased glucocorticoid and angiotensin II production, all activate the ubiquitin–proteasome system (UPS) to degrade muscle proteins. The initial step in this process is activation of caspase-3 to cleave the myofibril into its components (actin, myosin, troponin, and tropomyosin). Caspase-3 is required because the UPS minimally degrades the myofibril but rapidly degrades its component proteins. Caspase-3 activity is easily detected because it leaves a characteristic 14kD actin fragment in muscle samples. Preliminary evidence from several experimental models of catabolic diseases, as well as from studies in patients, indicates that this fragment could be a useful biomarker because it correlates well with the degree of muscle degradation in dialysis patients and in other catabolic conditions.
Maintenance of protein stores in chronic kidney disease
In uremic patients, protein stores are frequently depressed when assessed by a low serum prealbumin and weight loss, which includes loss of muscle mass [1, 2]. In pediatric patients with chronic kidney disease (CKD), linear growth is impaired and muscle mass is reduced. Although these findings have been attributed to “malnutrition”, many uremic patients with muscle wasting have not developed the problems because of inadequate diet; instead, they have complications that induce a complex series of physiological and biochemical adaptations, resulting in protein catabolism [3, 4]. In children and adults with CKD, these complications include metabolic acidosis, insulin resistance, increased glucocorticoid production, high levels of angiotensin II (Ang II), and inflammation [5–8]. Many observational studies and mechanistic investigations have attempted to explain this loss of protein stores, and especially the loss of muscle mass. There are at least three conclusions from these studies. First, rates of protein synthesis in muscle are generally unchanged, whereas rates of protein degradation tend to increase [5, 9]. Second, the daily rates of protein turnover in cells are so high (3.5–4.5 g protein/kg per day) that even a small increase in protein degradation (and/or a decrease in protein synthesis) will cause marked protein depletion over time [10]. Third, the increase in muscle protein degradation in uremia and most other catabolic disease states is mostly due to programmed activation of the ubiquitin–proteasome system (UPS) [5, 11]. Therefore, to understand muscle wasting, one must understand the UPS.
The ATP-dependent, ubiquitin–proteasome system (UPS)
Over the past two decades, progress in understanding the action and regulation of the UPS has been at the center of attempts to understand the control of protein turnover. The UPS includes concerted actions of enzymes that link ubiquitin (Ub), a member of the heat-shock protein family, to protein substrates that are destined for degradation (Fig. 1). When a chain of at least four to five ubiquitins are linked to a protein, it is marked for degradation in a second step mediated by the proteasome [12, 13]. Specifically, the tagged protein will be recognized by the 26S proteasome, a very large multicatalytic protease complex that not only recognizes Ub-conjugated proteins but also removes Ub, unwinds the protein, and injects it into the central core of the 26S proteasome. Once inside this central, tube-like structure, the protein substrate is degraded into small peptides [14].
Fig. 1The ubiquitin–proteasome pathway of protein degradation. Ubiquitin (Ub) is conjugated to proteins destined for degradation by an ATP-dependent process that involves three enzymes (E1–E3). A chain of five Ub molecules attached to the protein substrate is recognized by the 26S proteasome, which removes Ub and digests the protein into peptides. The peptides are degraded to amino acids by peptidases in the cytoplasm or used in antigen presentation. (Reproduced with permission from [15])
Three enzymatic components are required to link Ub to proteins that are destined for degradation. There appears to be only one E1 (Ub-activating) enzyme and around 40 E2 (Ub-carrier or conjugating) proteins. The key enzyme, however, is the E3 enzyme, which accounts for the exquisite specificity of proteins to be degraded. There are at least a thousand E3 enzymes (Ub ligase), and each can recognize a specific protein substrate and catalyze the transfer of an activated Ub from the E2 carrier protein to the substrate protein [10, 15].
Since the initial reports that the UPS recognizes specific proteins and tags them for destruction, knowledge about proteolytic processes in the proteasome has exploded. Thousands of proteins have been recognized as being degraded by the UPS, and novel cellular functions are now known to be regulated by Ub conjugation. In terms of protein breakdown, the major functions of the pathway are:
Rapid removal of proteins Protein degradation is irreversible, and hence, destruction of a protein generally leads to a complete termination of cellular process mediated by the protein. Consequently, protein degradation is critical for the regulation of metabolism and cell turnover. The rapid degradation of specific proteins also permits cells (as well as the organism) to rapidly adapt to a change in physiological conditions (e.g. requiring a switch to glucose as an energy source involves converting protein stores into amino acids that can be used in gluconeogenesis).
Regulation of gene transcription Ub conjugation affects gene transcription because many transcription factors become conjugated to Ub, and transcription activators are degraded by the proteasome [16]. This process regulates transcriptional activity by removing “spent” activators and resetting a promoter for additional rounds of transcription [17]. Second, the ability of transcription factors to function varies with their location within the cell. For example, nuclear factor (NF)-κB, a proinflammatory transcriptional factor, is kept outside the nucleus because movement into the nucleus is blocked by its association with an inhibitory protein, IκB. Destruction of IκB, initiated by the IKK kinase and carried out by the UPS, frees NF-κB, which then translocates to the nucleus to stimulate gene transcription [18]. A clinical application of this function of the UPS has developed in oncology. Bortezomib (Velcade, PS-341), a proteasome inhibitor, has proven to be beneficial in patients with multiple myeloma and is currently in clinical trials for the treatment of other cancers [19, 20]. The proposed mechanism of action involves the ability of bortezomib to prevent the UPS-induced destruction of IκB, thereby blocking the activation of NF-κB (an antiapoptosis transcription factor), leading to an increase in apoptosis [21]. Inhibition of the proteasome, therefore, will induce apoptosis of the neoplastic cells [22]. In addition, myeloma cells are also particularly dependent upon NF-κB to produce essential growth factors [especially inerleukin 6 (Il-6)]; when NF-κB is inactive, the growth of myeloma cells is depressed.
Quality-control mechanism The UPS selectively eliminates abnormally folded or damaged proteins that have arisen because of missense or nonsense mutations, biosynthetic errors, proteins damaged by oxygen radicals, or by denaturation. For example, in cystic fibrosis, the mutant form of the transmembrane conductance regulator protein (CFTR) is selectively degraded before it reaches the cell surface [23]. The UPS catalyzes destruction of this mutant CFTR because its tertiary structure is abnormal. Another example is the degradation of misfolded proteins within the endoplasmic reticulum. Endoplasmic-reticulum-associated degradation (ERAD) of proteins removes misfolded proteins by targeting them for destruction by proteasomes in the cytoplasm [24].
Influencing the function of the immune system The UPS is responsible for creating antigens from the degradation of foreign proteins (e.g. viral particles). The antigens are presented on the major histocompatibility complex as class I molecules. In this way, the 26S proteasome exerts dual roles of removing foreign proteins and creating a stimulus of the immune system [15, 25].
As a source of amino acids When carbohydrate calories are rapidly needed or when cells must respond to catabolic diseases/conditions, there is breakdown of cell proteins, especially skeletal muscle proteins. The UPS degrades muscle proteins to provide amino acids that can be converted to glucose (i.e. gluconeogenesis). An undesired consequence of this activity could be an inappropriate loss of muscle protein.
Functions of Ub not associated with proteolysis Ub can also be conjugated to proteins as a monomer (rather than as the typical Ub chain). When this occurs on cell-surface proteins, the protein is internalized into the endocytic pathway to be degraded in lysosomes [26, 27].
Uremia-activated mechanisms that accelerate loss of muscle protein
Results from rodent models of CKD have established that accelerated muscle protein catabolism involves many of the same cellular mechanisms that cause muscle wasting in other catabolic conditions, such as cancer, starvation, insulin deficiency/resistance, or sepsis [10, 28]. The principal mechanism causing muscle atrophy in CKD involves activation of the UPS. Evidence for this includes the presence of higher levels of mRNAs encoding certain components of the UPS, as well as a similar pattern of increases and decreases in the expression of about 100 atrophy-related genes (also called atrogenes) [5, 28]. Changes in atrogenes include decreased expression of various growth-related genes and increased expression of components of the UPS. Patients with different clinical conditions associated with muscle atrophy exhibit similar increases in mRNAs encoding components of the UPS (e.g. an increase in mRNAs encoding Ub and proteasome subunits) [11, 29–31]. In these cases, changes in gene expression are most likely due to transcriptional regulation because we have shown that uremia or abnormal insulin responses increase the transcription of Ub and subunits of the proteasome [5, 6, 32]. Additional evidence linking the UPS to protein degradation in catabolism is the finding that the increase in protein degradation in the muscle of rats with CKD (and other muscle-wasting conditions) can be blocked by inhibitors of the proteasome [5, 6, 33]. Considered together, these results indicate that muscle wasting is a specific and carefully orchestrated program.
Other questions are why are proteins degraded, and how is the complex program triggered in widely varied pathological conditions (e.g. acidosis in renal failure, low insulin levels in fasting and diabetes, inactivity, or glucocorticoids and cytokines in sepsis and other inflammatory conditions)? In fasting and in other disease states, acceleration of muscle-protein breakdown mobilizes amino acids, which are used for protein synthesis in tissues and for conversion to glucose in the liver [10]. However, if excessive protein degradation persists, the protein loss will have deleterious effects. In CKD, the breakdown of tissue protein produces nitrogenous waste products, which must be excreted to prevent the accumulation of uremic toxins [3, 10]. Finally, in muscle wasting conditions, contractile proteins are lost differentially, whereas in conditions causing atrophy (e.g. aging), all components of muscle cells seem to be affected.
What accounts for muscle-specific response? The answer to this question lies in the involvement of the UPS. The UPS degrades a specific protein depending on which E3 ubiquitin ligase is activated. For example, two Ub ligases, atrogin-1 (also known as MAFbx) and MuRF-1, are found specifically in muscle, and their expression increases dramatically (8- to 20-fold) in catabolic states, causing loss of muscle protein [28, 34, 35]. In mice lacking the genes for either atrogin-1 or MuRF-1, muscles grow normally, but in response to muscle denervation, the ensuing atrophy is 30–50% slower [34]. In addition, the muscle’s content of atrogin-1 mRNA can be considered a biomarker for the rate of proteolysis in muscles responding to a catabolic condition [36–38].
In uremia, the initial cleavage of myofibrillar proteins is mediated by caspase-3
Myofibrillar proteins (including actomyosin) comprise about two thirds of the protein in muscle, the major store of amino acids for new protein synthesis and for gluconeogenesis. The UPS readily degrades the main proteins in myofibrils (i.e. actin, myosin, troponin, or tropomyosin), but it does not readily break up the myofibril into its main component proteins [39]. This means that another proteolytic system must initially digest myofibrils to create substrates that can be degraded by the UPS [40].
Many catabolic conditions are associated with inflammation and/or cell injury, and these conditions activate a cysteine protease called caspase-3. We tested caspase-3 in an in vitro system using purified actomyosin and found that caspase-3 cleaves actomyosin and leaves a characteristic 14kD actin fragment [40]. When we activated caspase-3 in cultured muscle cells, we found that UPS rapidly degraded myofibrillar component proteins, and again, the 14kD C-terminal fragment of actin was left in the muscle cells [40]. The protein-cleaving action of caspase-3 is important because blocking caspase-3 will reduce overall protein degradation in muscle [40]. In addition, we have found the same cleavage processes are present in muscles of rodent models of uremia, type-1 or type-2 diabetes, or in Ang-II-induced hypertension [7, 35, 40, 41]. Moreover, we found that the 14kD actin fragment also accumulates in muscles of patients with loss of muscle mass due to painful osteodystrophy, uremia, or burn injury [42]. In the latter study, we found that the rate of protein degradation in muscle (measured from the turnover of labeled amino acids) directly correlated (r = 0.78) with the level of the 14kD actin fragment in the same muscle. In addition, there was a lower level of the 14kD actin fragment in muscle of hemodialysis patients who participated in 18 weeks of an endurance exercise training program. In summary, the level of the 14kD actin fragment is directly associated with measured protein degradation in muscle, and the accumulation of the fragment responds to a beneficial therapeutic intervention. If these properties hold up in other trials, the level of the 14kD actin fragment could be used as a “biomarker” of increased muscle protein degradation [42].
Signals triggering muscle atrophy in kidney disease and other catabolic states
Complications of CKD, as well as the complex syndrome of uremia, can trigger muscle protein breakdown. The triggering complications include metabolic acidosis, decreased insulin action (including insulin resistance), increased glucocorticoid production, high levels of Ang II, and inflammation [5–8]. Metabolic acidosis is known to cause accelerated protein degradation in infants, children, adults, the elderly, and patients with CKD (Table 1). The mechanism by which metabolic acidosis causes muscle wasting involves activation of the UPS and caspase-3 [40, 43]. In addition, acidosis changes hormone actions, such as insulin resistance and increased glucocorticoid production, which are involved in activating protein degradation [32, 41, 43–45] (Table 2). It is important to emphasize that the correction of metabolic acidosis has been shown to decrease protein breakdown in various clinical trials, indicating why maintaining normal serum bicarbonate levels should be part of standard clinical care (Table 1).
Table 1Evidence that metabolic acidosis induces catabolism of protein and amino acids in normal infants, children, and adults, as well as in patients with chronic kidney disease (CKD)Subjects investigatedOutcome measurementsTrial outcomeInfants [68]Low-birth-weight, acidotic infants were given NaHCO3 or NaClNaHCO3 supplement improved growthChildren with CKD [69]Measured rates of protein degradation in children with CKDProtein loss was ∼ 2-fold higher when HCO3 was < 16 mM compared with > 22.6 mMNormal adults [70]Acidosis induced and then measured amino acid and protein metabolismAcidosis increased amino acid and protein degradationNormal adults [71]Induced acidosis and then measured nitrogen balance and albumin synthesisAcidosis induced negative nitrogen balance and suppressed albumin synthesisChronic renal failure [72]Nitrogen balance before and after treatment of acidosisNaHCO3 improved nitrogen balanceChronic renal failure [73]Essential amino acid and protein degradation before and after treatment of acidosisNaHCO3 suppressed amino acid and protein degradationChronic renal failure [74]Muscle protein degradation and degree of acidosisProteolysis was proportional to acidosis and blood cortisolChronic renal failure [75]Nitrogen balance before and after treatment of acidosisNaHCO3 reduced urea production and improved nitrogen balanceHemodialysis [76]Protein degradation before and after treatment of acidosisNaHCO3 decreased protein degradationHemodialysis [77]Serum albumin before and after treatment of acidosisNaHCO3 increased serum albuminCAPD [78]Protein degradation before and after treatment of acidosisNaHCO3 decreased protein degradationCAPD [79]Weight and muscle gain before and after treatment of acidosisRaising dialysis buffer increased weight and muscle massCAPD continuous ambulatory peritoneal dialysisTable 2Metabolic acidosis in otherwise normal humans changed hormonal levels or responses to hormonesHormoneAcidosis-induced responseGrowth hormone (GH) [80–84]Suppressed GH secretionLower IGF-1 responseInsulin [44, 85, 86]Suppressed insulin-stimulated glucose metabolismInsulin-like growth factor (IGF)-1 [81, 84, 87]Decreased IGF-1 in plasma, and kidney and liver (but not in muscle)Thyroid hormone [82, 88]Decreased plasma T3 and T4 levels plus a higher plasma thyroid-stimulating hormoneGlucocorticoids [89]Increased glucocorticoid productionParathyroid hormone (PTH) [90, 91]Decreased sensitivity of PTH secretion to changes in plasma calciumVitamin D [91]Suppressed activation to 1,25 (OH)2 cholecalciferol
The finding that various diseases with muscle wasting are caused by activation of the UPS, plus the fact that coordinated changes in the expression of genes in muscle occur in different catabolic states, suggest that catabolic states activate a common cellular signaling pathway [28]. One signaling pathway is a decrease in phosphatidylinositol 3-kinase (PI3K) activity (Fig. 2). The involvement of this signaling pathway follows from the finding that several catabolic illnesses, including sepsis, acidosis, uremia, and diabetes, are characterized by insulin resistance [41, 44, 46, 47]. In normal muscle, binding of insulin or insulin-like growth factor (IGF)-1 to their receptors increases the activities of PI3K and its downstream target, Akt. In insulin-resistant conditions or with deficiency of IGF-1, the activity of this signaling pathway is depressed [35, 48, 49]. When PI3K activity falls, there is decreased production of phosphatidylinositol-3,4,5 phosphate (PIP3), leading to decreased phosphorylation and activity of the downstream serine/threonine kinase, Akt. This is a key step, because activated Akt is a major stimulator of growth-related processes via phosphorylation of the downstream kinases, GSK1 and mTOR/S6kinase, stimulating protein synthesis. On the other hand, reduced PI3K-Akt signaling (as occurs in insulin resistance) enhances protein breakdown in muscle [41]. The rise in muscle protein losses is associated with two catalytic processes: first, caspase-3 is activated to break down the complex structure of muscle; second, there is increased expression of the E3 ubiquitin ligases, atrogin-1 and MuRF-1, to degrade the proteins made available by caspase-3 [35–37] (Fig. 2). The result is muscle wasting.
Fig. 2The balance between muscle hypertrophy and atrophy depends on whether protein synthesis is more active than degradation or vice versa. In protein synthesis, insulin-like growth factor (IGF)-1 and insulin signaling leads to increased phosphatidylinositol 3-kinase (PI3K), which promotes Akt phosphorylation (Akt-P). Akt-P promotes phosphorylation of GSK1 and mTOR/S6 kinases, leading to increased protein synthesis. Akt-P also phosphorylates the forkhead (FoxO) transcription factor, preventing it from entering the nucleus to promote expression of atrogin-1, MuRF-1, and other atrogenes, thereby blocking protein degradation. In protein degradation, the opposite pathway happens, but additionally, decreased Akt-P leads to increased caspase-3 activity, further promoting degradation. In inflammation, it is thought that tumor necrosis factor (TNF)-α and other inflammatory cytokines phosphorylate the inhibitor of nuclear factor (NF)-κB (IκB), to free NF-B to enter the nucleus and promote MuRF-1 expression, and ultimately, muscle protein degradation. (Reproduced with permission from [15])
How are the two mechanisms activated? In studies of muscles from insulin-deficient rats or db/db mice (a model of insulin resistance), we found that accelerated muscle protein degradation increases the level of the proapoptotic factor, Bax [35, 41]. This is relevant because activated Bax causes the release of cytochrome C from mitochondria. Cytochrome C release, in turn, activates caspase-3 to trigger actomyosin/myofibril cleavage, leaving behind the 14kD actin fragment marker [35]. The activation of atrogin-1, and hence the UPS, involves another mechanism: changes in activity of the fork-head transcription factors (FoxO 1, 3 and 4). When these transcription factors are phosphorylated by Akt, they cannot enter the nucleus to stimulate transcription of atrogin-1. However, when PI3K/Akt activities are low, the FoxOs are not phosphorylated, so they can enter the nucleus to increase the transcription of atrogin-1, resulting in an increase in muscle protein degradation [35–37].
The influence of Akt on the other E3 ubiquitin ligase involved in muscle proteolysis, MuRF-1, is not as clearly established, and it may be linked to inflammation, as activation of NF-κB will cause overproduction of MuRF1 and muscle atrophy [37].
Besides acidosis and depressed insulin/IGF-1 action, another complication of CKD that participates in muscle wasting is increased glucocorticoid production. The complexity of these interactions is great because increased glucocorticoids can cause insulin insensitivity [32, 41, 50], and both insulin deficiency and insulin resistance increase glucocorticoid production. Glucocorticoids exert a permissive effect on protein degradation in muscle caused by several catabolic conditions. For example, activation of muscle protein breakdown does not occur in adrenalectomized animals with metabolic acidosis or with acute diabetes unless the animals are also given a physiological dose of glucocorticoids [32, 51–53]. Similarly, the increase in muscle proteolysis induced by Ang II or sepsis is largely eliminated by inhibiting the glucocorticoid receptor [7, 54]. This response to glucocorticoids is “permissive” because the same physiological level of glucocorticoids does not stimulate muscle protein degradation unless the animals are either acidotic or insulin deficient. These complex interactions actually make “physiological sense” because glucocorticoids evolved to integrate stress responses in different tissues. When glucose is needed, glucocorticoids mobilize amino acids from muscle protein. At the same time, glucocorticoids induce gluconeogenic enzymes in liver to catalyze the conversion of the amino acids to glucose and urea.
In children, accelerated loss of protein stores associated with glucocorticoid therapy frequently results in impaired linear growth. One glucocorticoid-dependent mechanism causing these defects is impaired response to insulin (insulin insensitivity). An allied mechanism is impaired response to IGF-1 arising from decreased or impaired action of growth hormone. Indeed, administration of growth hormone has been shown to improve linear growth of children being treated chronically with glucocorticoids [55]. Moreover, growth hormone has been shown to improve growth of children with chronic catabolic diseases such as CKD; whereas growth hormone improves growth, it does not improve growth in children to (or close to) a normal height for age [56–58]. The impaired response is likely due to the multiple complications of CKD, resulting in retarded growth. Interestingly, there is a small uncontrolled study of five children who were being treated with chronic intermittent hemodialysis and growth hormone. The children were then placed on an intensified daily hemodialysis regimen (3 h/day, five to six times a week) for a median time of 1.5 years [59]. The new regimen led to significant catchup growth into the range of a normal height for age. Intensified dialysis plus growth hormone could correct acidosis, improve insulin and IGF-1 signaling and responses to other hormones, and remove unidentified uremic toxins. These responses emphasize the complexity of sorting out mechanisms for muscle wasting in CKD.
Another catabolic factor associated with CKD is Ang II. Infusion of Ang II into rodents causes both anorexia and muscle protein loss by mechanisms that depend on glucocorticoids [7]. Also, there is the knotty problem of understanding the influence of inflammation. In this case, the mechanism(s) causing muscle wasting in inflammation is not clear. Suggested responses include the ability of certain inflammatory mediators to cause insulin resistance, as well as a more direct influence of inflammatory mediators on muscle protein metabolism (Fig. 2). The problem is difficult because the link between an increase in inflammatory markers in uremic patients (e.g. C-reactive protein) to loss of muscle mass has not been established [60, 61].
Finally, there is an intriguing protein—myostatin—a member of the transforming growth factor (TGF)-β family of cytokines. It is produced in skeletal and cardiac muscle and regulates muscle growth by limiting it [62]. Overexpression of myostatin in muscle leads to loss of protein mass via inhibition of Akt phosphorylation with an increase in active FoxO1; this increases the expression of atrophy-related genes [63, 64]. Underexpression of myostatin results in skeletal muscle hypertrophy [65]. An increase in myostatin expression is found in several cachexia-associated disease states. However, there is limited information about the influence of kidney disease on myostatin expression and function beyond changes in myostatin mRNA [66, 67].
Conclusion
In this brief review of mechanisms causing muscle protein losses, we discussed how a complex series of biochemical reactions are coordinated to create a genetic program that degrades muscle proteins. We also identified an initial step in muscle proteolysis that leaves behind a biomarker in muscle, the 14kD actin fragment, resulting from caspase-3. We emphasized how the UPS causes muscle wasting in uremia, as well as the role UPS plays in the regulation of cellular functions, ranging from the control of the cell cycle to activities that promote cancer. Indeed, inhibitors of proteasome activity have emerged as novel chemotherapeutic agents. Involvement of the UPS in such a wide range of functions explains why the 2004 Nobel Prize in Chemistry was awarded to Avram Hershko, Aaron Ciechanover, and Irwin Rose for their discovery of Ub and its role in orchestrating cellular protein turnover (http://nobelprize.org/chemistry/laureates/2004/).

muscle wasting
chronic kidney disease (ckd)
caspase-3
14kd actin fragment
protein degradation
uremia
muscle atrophy
ubiquitin-proteasome system (ups)


Single-Pair FRET Microscopy Reveals Mononucleosome Dynamics

We applied spFRET microscopy for direct observation of intranucleosomal DNA dynamics. Mononucleosomes, reconstituted with DNA containing a FRET pair at the dyad axis and exit of the nucleosome core particle, were immobilized through a 30 bp DNA tether on a polyethyleneglycol functionalized slide and visualized using Total Internal Reflection Fluorescence microscopy. FRET efficiency time-traces revealed two types of dynamics: acceptor blinking and intramolecular rearrangements. Both Cy5 and ATTO647N acceptor dyes showed severe blinking in a deoxygenated buffer in the presence of 2% βME. Replacing the triplet quencher βME with 1 mM Trolox eliminated most blinking effects. After suppression of blinking three subpopulations were observed: 90% appeared as dissociated complexes; the remaining 10% featured an average FRET efficiency in agreement with intact nucleosomes. In 97% of these intact nucleosomes no significant changes in FRET efficiency were observed in the experimentally accessible time window ranging from 10 ms to 10’s of seconds. However, 3% of the intact nucleosomes showed intervals with reduced FRET efficiency, clearly distinct from blinking, with a lifetime of 120 ms. These fluctuations can unambiguously be attributed to DNA breathing. Our findings illustrate not only the merits but also typical caveats encountered in single-molecule FRET studies on complex biological systems.
Introduction
Fluorescence (or Förster) Resonance Energy Transfer (FRET) is a process in which the energy of an excited donor fluorophore is transferred non-radiatively to an acceptor molecule [1]. The efficiency of energy transfer E is given by:
where R is the distance between donor and acceptor and R0 is the Förster radius, at which 50% energy transfer occurs (typically 5 nm for Cy3–Cy5, a commonly used FRET pair). FRET is a powerful tool to study the structure and function of biological molecules, such as DNA. When extended to the single-molecule level, single pair FRET (spFRET) can potentially be applied to determine the conformational distribution of an ensemble of molecules and the dynamics of individual molecules [2–4]. We exploited spFRET to study the structure and dynamics of single nucleosomes, the fundamental units of compaction and organization of eukaryotic DNA.
The nucleosome core particle consists of ∼ 50 nm DNA wrapped nearly twice around a histone-octamer protein-core [5]. Nucleosomal DNA has to unwrap from the nucleosome core to sterically allow processes such as transcription, replication and repair. Accessibility to nucleosomal DNA is facilitated by ATP-dependent remodeling enzymes in vivo [6]. However, it is known that spontaneous conformational changes of the nucleosome expose occluded sites in the DNA as well [7]. DNA breathing, the transient unwrapping and rewrapping of a stretch of DNA from the nucleosome core, has recently been studied in detail with a variety of fluorescence techniques. The equilibrium constant of this process was determined with bulk FRET measurements [8]. Unwrapping lifetimes of 10–50 ms were obtained with stopped-flow FRET measurements and Fluorescence Cross Correlation Spectroscopy (FCCS) [9]. Interestingly, based on their single pair FRET work, Tomschik et al. concluded that unwrapping of nucleosomal DNA occurs to a much larger extent than was previously anticipated [10]: they suggested that 30–60% of the nucleosomal DNA was unwrapped with a lifetime on the order of ∼150 ms before rewrapping.
Although the conceptual beauty of FRET studies is undisputed, there are a number of important caveats in single-molecule FRET studies of biomolecules, such as fluorophore blinking, photobleaching and sample immobilization. Here, we addressed these issues. spFRET microscopy on mononucleosomes revealed two dominant types of dynamics: acceptor blinking and intramolecular rearrangements that we attribute to DNA breathing, which only became apparent after suppression of blinking. Upon immobilization, we observed three different populations: 90% of the nucleosomes dissociated or represented donor-only species, and 10% remained intact. Of these fully wrapped nucleosomes, 97% showed stable FRET on timescales between 0.01–10 s, while 3% showed dynamics with a dwell time of 120 ms that we attribute to conformational changes in the nucleosome.
Material and methods
DNA preparation
A 177 base pair (bp) DNA was constructed by PCR using the 601 nucleosome positioning element [11] as template. PCR primers were as follows. Forward primer: Biotin-TTTGAATTCC CAGGGAATTG GGCGGCCGCC CTGGAGAATC CCGGTGCCGA GGCCGC (acceptor labeled nucleotide is underlined). Reverse primer: ACAGGATGTA TATATCTGAC ACGTGCCTGG AGACTAGGGA GTAATCCCCT TGGCGGTTAA AACGCGGGGG ACAGCGCGTA CG (donor labeled nucleotide is underlined). We used either Cy3–Cy5 or ATTO550-ATTO647N as donor-acceptor FRET pair. PCR products were purified with a GFX PCR DNA & Gel Band Purification Kit (GE Healthcare). The position of the labels was chosen such that after reconstitution the acceptor was located at the nucleosome exit, and the donor near the dyad axis, as illustrated in Fig. 1. Donor and acceptor were predicted to be ∼4 nm apart, as deduced from the nucleosome crystal structure [12], resulting in a FRET efficiency E of approximately 0.8 for the Cy3–Cy5 pair (R0 ∼5 nm), and of approximately 0.9 for the ATTO550-ATTO647N pair (R0 ∼6 nm).
Fig. 1FRET system for the study of mononucleosome dynamics. a The 177 bp DNA construct, indicating the position of the labels 80 bp apart in a fragment containing the 601 nucleosome positioning sequence. A biotin label allowed for immobilization of the construct. b, c Illustrations of the mononucleosome structure, indicating the position of donor and acceptor upon reconstitution. The distance between the labels was ∼4 nm, at which efficient FRET takes place. Unwrapping of the DNA from the nucleosome core will be accompanied by a decrease in FRET due to increasing separation between donor and acceptor
Nucleosome reconstitution
Recombinant histone octamers were mixed with the DNA construct at a 1:1 ratio, in TE (1 mM EDTA, 10 mM TRIS pH 8.0) and 2 M NaCl. Mononucleosomes were reconstituted by salt dialysis against 0.85, 0.65, 0.5 and finally against 0.1 M NaCl, all buffered with TE.
Bulk fluorescence measurements
Bulk fluorescence experiments were carried out on a Luminescence Spectrometer (LS55, Perkins Elmer). All experiments were performed at room temperature (22°C). The nucleosome concentration was 10–50 nM. The donor dye was excited at 515 nm and the emission was recorded from 535 to 700 nm. The acceptor dye was excited at 615 nm and the emission was recorded from 635 to 700 nm, to obtain acceptor-only emission spectra. The FRET efficiency was determined from the enhanced fluorescence of the acceptor using the (ratio)A method [13]:
where  and  are the acceptor and donor extinction coefficient respectively at wavelength λ,  is the fluorescence intensity of the acceptor when excited at wavelength λ, and d+ is the fractional labeling coefficient of the donor. The fluorescence intensity of the acceptor was determined at its maximum value. d+ was determined from DNA and fluorophore absorption peaks in an absorption spectrum of the labeled DNA, measured from 230 to 700 nm with a spectrophotometer (Pharmaspec UV-1700, Shimadzu).
Single-molecule FRET measurements
Cleaned glass slides were amino functionalized with 10 μg/ml poly-d-lysine, and subsequently incubated for 4 h with an amine reactive polyethylene glycol (PEG) mixture: 20% mPEG-succinimidyl propionate 5,000 molecular weight (Nektar Therapeutics) and 0.2% biotin-PEG-n-hydroxysuccinimide 3,400 molecular weight (Nektar Therapeutics) in 0.1 M sodium carbonate buffer (pH 8.2). A flow cell was assembled by sealing a poly-dimethylsiloxane channel with a PEG functionalized slide. A 0.1 mg/ml streptavidin (Roche) solution was incubated for 5 min, and subsequently washed away. A sample, which typically consists of 10–50 pM of labeled mononucleosomes in 50–200 mM NaCl, 10 mM TRIS.HCl pH 8.0, 0.03% NP-40, and 10–50 nM unlabeled mononucleosomes, was injected in the channel and immobilized. An enzymatic oxygen scavenger system (1% glucose, 2% β-mercaptoethanol (βME) or 1 to 2 mM Trolox (Sigma), 0.2 mg/ml glucose oxidase, and 0.04 mg/ml catalase), was added to the buffer to extend the lifetime of the fluorophores before photobleaching. The buffer was degassed prior to use to further reduce the oxygen concentration. The flow cell was mounted on a microscope equipped with a 100X oil-immersion TIRF microscope objective (NA = 1.45, NIKON) and temperature-stabilized at 22°C using a water circulating bath connected to all parts of the setup in contact with the sample. The 514 nm line of an Ar+ laser (Coherent) was used to illuminate an area of ∼ 600 μm2 with a power of 0.9 mW. In the case of alternating excitation, a 636 nm diode laser (Power Technology) was used to illuminate an area of ∼900 μm2 with a power of 0.3 mW. Both beams were circularly polarized and were displaced parallel to the optical axis of the objective, so that an evanescent excitation field was generated by total internal reflection of the light at the glass-water interface. The excitation intensity at the interface in the evanescent field is ∼4 times higher than the incident beam intensity at the critical angle [14]. We therefore estimated that the resulting excitation intensities at the interface were ∼0.6 kW/cm2 for the 514 nm excitation and ∼0.13 kW/cm2 for the 636 nm excitation respectively.
The fluorescence was collected by the objective and filtered through a custom-made dual color band pass filter (Chroma), that rejects scattered laser light, and a long pass filter (OG530, Schott). The fluorescence was further split into a donor and an acceptor channel by a custom-made dichroic wedge mirror (0.5° angle, center wavelength of 630 nm, Chroma) placed in the infinity path of the microscope [15]. A +150 mm achromatic lens (Thorlabs) projected the separate images on a multiplication gain CCD camera (Cascade 512B, Roper Scientific) operating at a frame rate of 20 to 100 Hz.
Data analysis
The simultaneously acquired donor and acceptor images (typically 80 by 80 pixels) were aligned with respect to one another through their cross correlation. The first 50 donor and acceptor frames were overlaid, and their intensities averaged. Low frequency background signal was filtered out with a high-pass FFT filter. The location of the fluorophores was then determined by applying a threshold of 2 times the background noise level. A time-trace of donor and acceptor intensities was then calculated by integrating the pixel intensities 1.5 pixel around the fluorophore center for each frame and each image. In the case of alternating excitation, the acceptor intensity upon direct excitation was retrieved by deinterleaving the acceptor time-trace. The FRET efficiency was calculated from [2]:
where IA and ID are acceptor and donor intensity respectively, and  is a parameter to correct for photophysical properties of the dyes. ΦA and ΦD are acceptor and donor quantum yield, and ηA and ηD are acceptor and donor detector efficiency respectively. As a first approximation γ was set to unity.
A more accurate estimate for γ was obtained from experimental intensity time traces where donor bleaching took place after acceptor bleaching. In these cases the FRET efficiency could also be calculated from donor quenching:
where ID0 is the donor intensity after bleaching of the acceptor. Combining Eqs. 3 and 4 results in:
Experimental results
Bulk fluorescence spectra reveal proper reconstitution of mononucleosomes
The results of bulk fluorescence and absorption experiments on reconstituted mononucleosomes are shown in Fig. 2. The reconstituted sample showed efficient FRET, indicated by a distinct peak of fluorescence at the acceptor maximum emission wavelength (670 nm for ATTO647N, see Fig. 2a). This peak was not present in the labeled DNA-only sample, confirming that the donor and the acceptor were in close proximity due to mononucleosome reconstitution. As a control, we diluted the mononucleosome sample in 2 M NaCl, as this high ionic strength disrupts nucleosome structure [16]. As predicted, over 90% of the energy transfer signal was lost.
Fig. 2Bulk fluorescence emission and absorption spectra revealed proper reconstitution of mononucleosomes. a Bulk fluorescence emission spectra. A distinct peak of fluorescence at the acceptor emission wavelength was seen after reconstitution, which was not present for the labeled DNA only. The peak disappeared upon dilution of the reconstituted material in 2M NaCl, an ionic strength at which nucleosome structure is disrupted. b Bulk absorption spectrum of the fluorescently labeled DNA construct. The stoichiometry of the labels was obtained by comparison with the absorption spectra of ATTO550 and ATTO647N (as provided by the manufacturer), which are plotted with dotted lines
The observed average FRET efficiency in the reconstituted mononucleosomes was 0.75 ± 0.1, which was in good agreement with FRET values predicted by the position of the FRET pair in the nucleosome. From the bulk FRET experiments, and the predicted maximum FRET efficiency of ∼0.9 for a mononucleosome with the ATTO550-ATTO647N FRET-pair, we estimated the reconstitution yield to be at least 85%. Some residual donor emission can be accounted for by incomplete acceptor labeling. With absorption measurements on the DNA construct (Fig. 2b) we determined that the acceptor:donor:DNA stoichiometry was ∼0.7:0.9:1. Together, these bulk data show that the labeled DNA construct and the histone proteins properly formed mononucleosomes upon reconstitution (see Fig. 1).
spFRET microscopy reveals individual nucleosomes together with a large population of dissociated nucleosomes
To investigate mononucleosome subpopulations and dynamics, spFRET measurements were performed in a wide field microscope. Figure 3 shows an example of typical single-molecule fluorescence images of immobilized mononucleosomes. In Fig. 3a, an acceptor channel image was superimposed on a donor channel image. 10% of the immobilized fluorophores showed efficient FRET, as indicated by colocalized fluorescent spots in the acceptor channel upon donor excitation, and thus represent fully reconstituted mononucleosomes.
Fig. 3Single molecule fluorescence image of immobilized mononucleosomes. a False color representation of averaged donor and acceptor channel images, excited at 514 nm. The arrows point at molecules that featured efficient FRET from donor to acceptor. The majority of the molecules however did not show FRET and appears in red. b The same field of view excited at 636 nm, allowing for unambiguous identification of acceptor fluorophores
In contrast, 90% of the fluorophores did not show FRET at all. This conflicts with the bulk experiments, where after correction for incomplete labeling an average FRET efficiency of 0.75 was found. As mentioned before, there was a fraction of donor only labeled species (∼30%), but this alone could not explain the observed discrepancy between bulk and single-molecule measurement. When we directly excited the acceptor fluorophores (see Fig. 3b), we found that most of the donors were colocalized with an acceptor. Therefore we conclude that FRET signal was lost during the single-molecule measurement, due to disassembly of a large fraction of the nucleosomes.
It is known that nucleosomes become unstable and dissociate when they are diluted to low concentrations [17, 18]. For our wide field spFRET measurements we diluted to pM fluorophore concentrations to resolve individual fluorophores. We ensured that the nucleosome concentration was always above 10–50 nM. By adding an excess of unlabeled mononucleosomes and 0.03% non-ionic detergent (NP-40) to our buffer. Thastrom et al. [19] reported that under these conditions nucleosomes do not dissociate in bulk solutions. We found that even 50 nM of unlabeled nucleosomes, far above the dilution-driven dissociation threshold, did not retain proper nucleosome folding, excluding dilution effects to be the cause.
It is known that H2A–H2B histone dimers can spontaneously be exchanged from the protein core [20], which in our case would result in a transient loss of FRET. However, we found the same amount of disassembled nucleosomes upon immobilization when the octamer protein core was crosslinked by dialysis against 0.05% glutaraldehyde in 1 mM EDTA. We confirmed that the crosslinking itself did not dissociate nucleosomes with bulk fluorescence experiments. This suggests that not the histone protein core dissociates, but rather that the wrapped DNA loosens or significantly rearranges itself around the protein core. We confirmed that mononucleosomes in free solution (in the same buffer used for single-molecule experiments) remain stable for hours at room temperature using bulk fluorescence measurements. Therefore we consider the dissociation of the nucleosomes described here to be associated with their immobilization to the functionalized cover glass. As an alternative immobilization strategy we performed experiments with biotinylated BSA-functionalized cover glasses instead of PEGs. Biotinylated BSA is often used for single-molecule studies involving nucleic acids, whereas PEGs are often used for studies involving DNA-protein complexes [21]. Biotinylated BSA yielded even less intact nucleosomes. The exact nature of the interactions of the nucleosomes with the modified cover slides remains unclear, but the destabilizing effect of the surface forms a hurdle for obtaining large datasets of spFRET measurements.
On the 10% immobilized mononucleosomes showing FRET, irreversible loss of FRET was only found after photobleaching, implying that their nucleosomal structure remained intact after immobilization.
Single-molecule fluorescence footprint of individual nucleosomes
Example intensity time-traces of intact single nucleosomes are shown in Fig. 4a and b. Donor and acceptor intensity were clearly anti-correlated, indicative of their FRET interaction. The intensity of a single donor (Cy3) was 1.4 ± 0.3 × 103 counts/10 ms at a signal to noise ratio (SNR) of 5. When the donor was quenched by FRET, both the intensity and SNR decreased. The intensity of an acceptor (Cy5) excited via FRET was 0.9 ± 0.2 × 103 counts/10 ms at a SNR of 3. After ∼4 s of continuous illumination at ∼0.6 kW/cm2 either donor or acceptor photobleached, limiting the observational window to a few seconds. The total number of emitted photons until bleaching from a FRET pair was ∼105 (calculated with gain G = 33 counts/photon, detection efficiency ηD and ηA ∼15%). The observed average FRET efficiency of the high FRET level was ∼0.5 ± 0.13, slightly lower than the values measured in the bulk. From the traces where donor bleaching takes place after acceptor bleaching, we estimated the correction factor ã for photophysical parameters of Cy3/Cy5 to be ∼0.7 ± 0.3 (Eq. 4). The corrected FRET efficiency was then ∼0.6 ± 0.3, in good agreement with values obtained from bulk measurements.
Fig. 4Single molecule FRET traces from individual mononucleosomes. The top panels in a and b show the intensity time traces of donor and acceptor for green excitation; the middle panel shows the intensity time traces of donor and acceptor for red excitation, which were acquired in alternation with the green excitations. The bottom panels show the calculated FRET efficiency. The fluctuations between high and low FRET states featured perfect correlation with the corresponding acceptor intensity traces excited at 636 nm. c Histogram of the FRET efficiencies of multiple single molecule traces. d Histogram of the FRET efficiencies of a single trace. The spread in FRET efficiency was larger between different traces than within a single trace
We found that the standard deviation in FRET efficiency of the high FRET state of the entire population (0.13) was larger than the standard deviation within individual traces (0.06), as shown in the histograms in Fig. 4c and d. This observation can be accounted for either by different nucleosome populations with slight variations in FRET efficiency, or by local differences in rotational freedom of the dyes due to immobilization.
Acceptor blinking is the dominant source of spFRET dynamics
The FRET traces shown in Fig. 4a and b are highly dynamic and fluctuate between a high FRET state (E ∼0.6, lifetime 2.5 s) and a low FRET state (E ∼0.1, lifetime 0.13 s). Interestingly, the characteristics of these fluctuations, i.e. both on and off time, and the low FRET level, are remarkably similar to those observed by Tomschik et al. [10] who performed analogous experiments. This similarity however, is remarkable in view of the completely different FRET-label location. Tomschik et al. labeled the nucleosome opposite to the dyad axis and probed the DNA at the most internal position of the nucleosome, whereas our substrate has labels at the most exterior position. Widom et al. have previously shown that the enzymatic accessibility of the DNA inside a nucleosome strongly reduces as the DNA is more internal in the nucleosome [7], suggesting a higher frequency of unwrapping events in our experiments.
Because of the nearly complete absence of acceptor emission, we investigated the nature of these fluctuations in order to exclude reversible transitions of the acceptor to an inactive state (acceptor blinking, resulting in a Förster radius of effectively zero [22]) as the origin of these events. By alternating donor excitation with direct acceptor excitation we could directly monitor the acceptor condition as shown in Fig. 3b. After deinterleaving the data into two time-traces, one for green excitation and one for red excitation, it became obvious that the fluorescence intensity of the acceptor upon direct excitation correlated perfectly with the enhanced emission of the acceptor due to FRET. Thus, the low FRET state must be attributed to blinking, due to a dark-state level of the acceptor. Further evidence that these fluctuations were caused by acceptor blinking was provided by experiments with alternative acceptor dye (ATTO647N, emission spectrum similar to Cy5). Alternating excitation of the acceptor dye revealed a strong positive correlation between sensitized emission of the acceptor and direct excitation of the acceptor. In this case the low FRET state was also present, but with a much shorter lifetime of 0.046 s. In conclusion, our data confirm that the fluctuations between a high and a low FRET state reflect photophysical processes in the acceptor dye rather than nucleosome conformational changes.
We further analyzed the single-molecule FRET traces for dynamics other than blinking. Therefore we filtered out blinking events by the application of a threshold on low FRET efficiencies (≤ ∼0.1–0.2, dependent on the noise in the measurement). Although careful inspection did occasionally reveal anticorrelated features of donor and acceptor channel, these features had a lifetime below the time resolution of our measurements. To confirm that we did not overlook any dynamics, we analyzed the fluorophore intensity noise in the high FRET state, which in the absence of dynamics should be limited by shot noise. The theoretical noise σtot in the measurement was estimated by [23]:
where G is the multiplication gain factor, F is the excess noise factor due to the multiplication gain register, S is the number of photons that reach the camera, Φ is the camera quantum yield, D is the dark count, and σR is the readout noise. The first contribution represents photon shot noise after multiplication, the second contribution represents the camera dark noise after multiplication, and the third the ADC converter electronic noise. Readout noise and dark noise were calculated from the standard deviation of an area of the chip that was not illuminated by fluorescence to be 130 counts/10 ms. The actual noise σ in the single-molecule fluorescence traces was estimated by the standard deviation of the measured fluorophore intensity. The measured and calculated noise were tested for equality with an F test:  where α is the significance level at which the test was performed (0.05), and ν1, ν2 are the degrees of freedom used to calculate σ and σtot respectively. We found that the total measured noise was significantly (typically 1.5 times) higher than that predicted by photon statistics and camera noise only. This implied that the traces contained dynamic events that cannot be fully resolved, originating from either photophysical processes (short blinking events, or intersystem crossing), or fast nucleosome dynamics. Hence, to accurately capture these events, blinking had to be further suppressed, and the sampling frequency had to be increased.
Suppression of blinking
In order to suppress blinking, we first tested a different acceptor dye (ATTO647N), which was reported to have superior photochemical stability compared to Cy5 [24]. As mentioned before, this acceptor dye showed blinking as well, as seen in the example traces and histograms of Fig. 5a and b. Although a small fraction of molecules did not show any dynamics in FRET, the majority significantly blinked. In the case of Cy5 93% of all acceptors excited via FRET showed significant blinking, with a lifetime of ∼0.13 ± 0.05 s, and lifetime of the high state of ∼0.8 ± 0.1 s. In the case of ATTO647N, 94% of all acceptors excited via FRET showed blinking, with a lifetime of ∼0.046 ± 0.02 s, and lifetime of the high state of ∼1.2 ± 0.2 s. In conclusion, the use of a different dye did not suppress blinking to the required level, but just yielded different blinking statistics.
Fig. 5Fluorophore blinking in spFRET traces obtained from mononucleosomes. In the presence of βME, nucleosomes labeled with Cy5 a or ATTO647N b both show severe blinking in 95% of the traces. Example traces (top) and blinking lifetime histograms (bottom) are shown. c,d In the presence of an alternative triplet quencher, Trolox, blinking of both dyes is significantly suppressed. The example traces (top) show the absence of blinking in 90% of the traces, while the blinking lifetime histograms (bottom) show a small but finite amount of fast blinking still present in ∼10% of the traces
Recently a different approach to reduce blinking was described by Rasnik et al. [25]. They replaced the triplet quencher βME in the oxygen scavenger system by a water-soluble analog of vitamin E, Trolox. Using this approach, Cy5 blinking in single-molecule FRET measurements on DNA constructs was eliminated. We tested the effect of Trolox in the imaging buffer on blinking of FRET pair labeled mononucleosomes. Results are shown in example traces and histograms in Fig. 5c and d. Both for Cy5 and ATTO647N blinking was dramatically suppressed in the presence of 1 to 2 mM Trolox: over 90% of the traces showed no observable blinking. Noise analyses of the intensity fluctuations in most of these traces were fully accounted for by camera noise and photon statistics (shot noise) only. Thus, within our time resolution (10 ms), no effect of short time scale blinking, or inter system crossing, was detected.
Surprisingly, less than 10% of the observed FRET pairs still showed some extent of blinking indicated by fast excursions into a FRET state below 0.2, with a typical off time of 14 ± 1 and 13 ± 1 ms respectively, as shown in the histograms in Fig. 5c and d. Because the lifetime of these blinking events was on the order of the smallest sampling time used, blinking events were not identified by alternating excitation of the acceptor dye, but only by FRET efficiencies below the noise threshold. Direct excitation of the acceptor did only reveal some occasional blinking in the acceptor traces, with the same lifetime of 13–14 ms. To confirm that a small but finite amount of fast blinking still occurred in the presence of Trolox, we performed spFRET measurements on a FRET pair that was separated by 11 basepair duplex DNA. This construct does not exhibit structural changes that affect the FRET intensity. In this case we also observed a small, but finite amount of blinking in a number of traces (data not shown), with a lifetime similar to that measured on mononucleosomes.
A fraction of the immobilized nucleosomes shows dynamics clearly distinct from blinking
The suppression of blinking finally allowed us to unambiguously identify non-blinking dynamic events in the FRET traces. From a sample of 236 mononucleosomes that showed FRET, we found that over 95% of the traces essentially show stable FRET efficiency, as illustrated in Fig. 6a and b; all anti-correlated features in the FRET efficiency were short-lived and fall within the noise of the measurement. Thus, the upper limit for dynamic events that could have been missed in this population was 10 ms (the sampling time used). Interestingly, 3% of the traces showed dynamic events clearly distinct from blinking (examples shown in Fig. 6c and d), as judged by the following criteria: (1) the acceptor signal of a low FRET event was significantly higher than zero. (2) No correlated change in acceptor intensity was detected using alternating excitation. (3) Events persisted for at least two data-points. We found 14 events with an average FRET change ΔE of −0.23 and an average dwell time of 120 ± 5 ms, as summarized in the histogram and cumulative distribution function in Fig. 6e and f respectively. The lifetime was determined by fitting a cumulative exponential distribution to the data, independent of binning and therefore a more accurate way of determining the lifetime when using small datasets (Fig. 6f) than fitting a distribution to binned data. The lifetime of the high FRET state could not be determined accurately, due to the short time window that was available due to photobleaching. Since we explicitly checked the vitality of both fluorophores, we ruled out photodynamics and we could unambiguously attribute the observed features to DNA breathing dynamics.
Fig. 6A fraction of the immobilized nucleosomes showed dynamics clearly distinct from blinking. a,b After suppression of blinking with Trolox, over 95% of the FRET traces do not show FRET dynamics. The theoretical photon and instrument noise is approximately indicated by the grey bars. c,d ∼3% of the intensity traces (top panels) showed FRET fluctuations (bottom panels) clearly distinct from blinking: the acceptor intensity was significantly higher than zero, and events persisted multiple data points (see insets). These fluctuations clearly exceeded the noise. e Histogram and cumulative distribution plot f of the lifetime of the dynamic events. An exponential fit to the data gave an average lifetime of 120 ms
Discussion and conclusion
Time-traces of spFRET microscopy on single reconstituted mononucleosomes revealed two types of dynamics: acceptor blinking and intramolecular rearrangements. Intramolecular rearrangements became only apparent after suppression of blinking. Both Cy5 and ATTO647N showed severe blinking in a deoxygenated buffer in the presence of 2% βME. Replacing the triplet quencher βME with Trolox effectively eliminated most blinking effects. The lifetime of DNA unwrapping that we obtained after rigorous elimination of blinking events (∼120 ms) was comparable to the 150–180 ms obtained by Tomschik et al. [10], despite the very different location of the labels in the nucleosome, and probably less important, the different DNA sequence and origin of the histones. However, we observed a very similar lifetime (∼130 ms) for Cy5 blinking under comparable buffer conditions (2% βME). The FRET efficiency of the open states in our experiments was significantly above the detection threshold, so we can explicitly exclude photophysics as the origin of the observed changes in FRET efficiency.
Our single-molecule measurements revealed at least three subpopulations in the reconstituted and immobilized nucleosome sample: 90% of the fluorophores represented dissociated nucleosomes or donor only species, 10% represented intact nucleosomes. Of these, 97% remained stable on time-scales ranging from 10 ms to 10 s of seconds, while 3% showed intervals with reduced FRET efficiency and a lifetime of 120 ms clearly distinct from blinking.
Why most nucleosomes dissociate upon immobilization to the cover slip remains unknown. Immobilization of the molecules is necessary for extension of the available observation time. The time limit is given by photobleaching, one of the key advantages of this method with respect to, for example, Fluorescence Correlation Spectroscopy. However, the close proximity to the surface provides ample opportunity for interactions with it. Surface induced nucleosome dissociation has been reported before in Atomic Force Microscopy (AFM) studies. Using AFM in liquid, Nikova et al. observed an unwrapping of ∼25 nm of DNA from nucleosomes absorbed to a mica surface [26]. This unwrapping was attributed to a depletion of H2A–H2B histone dimers induced by the high surface charge of the mica, resulting in unwrapping of DNA. Although PEGs are neutral polymers that are commonly used to reduce non-specific surface binding of proteins, they may affect nucleosomes in different ways: PEG molecules have been reported to interact strongly with unfolded proteins [21], and could therefore possibly interact with histone tails. Furthermore, histone proteins are known to be adhesive to glass or plastic [27].
The large fraction of dissociated nucleosomes we report here was not observed by Tomschik et al. [10]. Because of the internal position of the labels they used, at least 50 bp of DNA had to be detached from the histone core before FRET was completely lost. We labeled the DNA at the very end of the histone bound part, and accordingly a detachment of 10–20 bp of DNA would already result in complete loss of FRET. Furthermore, the exterior part of the DNA is largely constrained by the mobile H2A–H2B dimer, whereas the labeled part of the DNA in the nucleosomes used by Tomschik et al. is mostly constrained by the more stable H3–H4 tetramer.
A labeling strategy by Li et al. [8, 9], who end-labeled a 601 nucleosome positioning element together with either histone H3 or H2A, provides a more comparable construct. Based on stopped-flow FRET and FCCS experiments they deduced an unwrapping rate of 4 s−1 an unwrapping lifetime of 10–50 ms. The 3% of our traces that showed dynamics typically featured multiple unwrapping events before photobleaching. Though photobleaching obstructs quantification of the unwrapping rate, it is of the same order of magnitude as observed by Li et al. The lifetime of the unwrapped state we observed is five to ten times larger. This discrepancy may in part be explained by differences in experimental conditions and nucleosome constructs; we can however not exclude the possibility that we overlook short-lived unwrapped states, biasing our data to a longer lifetime.
The absence of observations of DNA unwrapping in the majority of the intact nucleosomes reported in this study is in strong contrast with the extent of DNA breathing dynamics found by Li et al. [8, 9]. Two possible explanations could account for this difference: (1) The most frequently occurring DNA unwrapping occurs at a rate that exceeds the time resolution of our experiment. The rare dynamics (3%) that we observe would reflect the release of multiple histone-DNA contacts, a process that would occur less often and on longer time scales than unwrapping of only the first DNA-octamer. However, unwrapping of 10–20 bp of DNA would induce a more dramatic reduction in FRET efficiency than the reduction we observed, which is consistent with unwrapping of 10 bp or less. (2) The immobilized nucleosomes did not undergo breathing dynamics. It should be kept in mind that because of the disruption of 90% of the nucleosome upon immobilization, we only probed a subset of nucleosomes that do not dissociate upon immobilization. These nucleosomes could either be resistant to unwrapping of the DNA, or immobilized in such a way that DNA dynamics are inhibited due to interactions with the surface, while still retaining proper folding. In either case, immobilization is expected to have major impact on nucleosome dynamics, emphasizing the need for a more inert immobilization than point attachment to a PEG coated surface.
Our findings demonstrate that experimental conditions can have a profound impact on the data obtained when probing nucleosome structure and conformational dynamics. Immobilization effects and blinking dynamics have to be accounted for, and where possible suppressed in order to extract biologically relevant data from spFRET experiments. We have shown that DNA breathing kinetics obtained from carefully optimized spFRET experiments approaches values obtained from bulk experiments, opening the way to more complex single-molecule studies of chromatin dynamics.

fret nucleosome blinking single-molecule