The clinical diagnostic groups did not differ by age, gender, years of education, or postmortem interval (**Table 1**). There were more subjects with an ApoE 4 allele in the aMCI (33%) and AD (44%) groups than in the NCI group (0%). AD cases had significantly lower MMSE scores compared to both aMCI and NCI cases (p < 0.001), whereas the latter two groups did not differ statistically (**Table 1**). GCS z-scores were significantly lower in AD compared to the NCI and aMCI groups (p < 0.0001). Subjects in the different clinical diagnostic groups displayed considerable heterogeneity with respect to pathological diagnostic criteria. Neuropathological examination revealed that 58% of NCI, 60% of aMCI, and 90% of AD cases were classified as Braak stages III–VI. Using NIA-Reagan criteria, 25% of NCI, 50% of aMCI, and 90% of AD cases were classified as intermediate to high likelihood of AD (**Table 1**). For CERAD diagnosis, 42% of NCI, 30% of aMCI, and 100% of AD cases received a diagnosis of probable/definite AD. Statistical analysis revealed that the AD group displayed significantly higher pathology than the NCI and aMCI groups based on Braak (p < 0.001), NIA-Reagan (p < 0.001) and CERAD (p = 0.008) diagnosis.

#### TABLE 2 | Top 100 miRNAs dysregulated in AD compared to NCI frontal cortex.

#### TABLE 2 | Continued

(Continued)

\*Star strand of miRNA.

### miRNA Expression Dynamics during the Progression of AD

A small pilot miRNA microarray screen performed on RNA isolated from frozen frontal cortex tissue from three NCI and AD cases showed that the expression levels of 30 miRNAs were significantly different between the two groups, along with miRNAs that showed high but non-significant magnitudes of expression change between the two groups. The top 100 miRNAs are listed in **Table 2**. We performed qPCR to measure select miRNAs in frontal cortex across all groups examined. Specifically, we tested the expression levels of 20 of the top 100 miRNAs based on significant differences, magnitude of expression level difference and/or interest based on a literature search (Cogswell et al., 2008; Hébert et al., 2013; Lau et al., 2013; Wong et al., 2013) (**Table 3**). Of the 20 candidate miRNAs evaluated, miR-498, and miR-150 were significantly upregulated in AD frontal cortex; miR-150 was up-regulated in aMCI. Several miRNAs were also significantly down-regulated in AD frontal cortex, including miR-886-3p, miR-132, miR-21, miR-23a, miR-140-3p, miR-212, miR-23b, let-7d, and miR-181a (**Table 3**). However, two distinct clusters of these miRNAs, miR212/132, and miR 23a/23b, were also significantly downregulated by ∼50% in the frontal cortex of aMCI subjects relative to NCI subjects (**Table 3**, **Figures 1A,B**). By contrast, qPCR analysis of temporal cortex revealed that miR-212 and miR-132 expression levels were decreased only in the AD group, whereas miR-23a and miR-23b were unchanged across the clinical diagnostic groups (**Table 4**). Since miRNAs negatively regulate transcript stability, down-regulation of the

\*p < 0.05, \*\*p < 0.01.

miR-212/132 and miR-23a/b clusters in frontal cortex would be predicted to promote the stability of their target mRNAs in aMCI. Given several lines of evidence that the frontal cortex undergoes neuroplastic, presumably neuroprotective remodeling in the face of mounting AD neuropathology during MCI (DeKosky et al., 2002; Counts et al., 2006; Bell et al., 2007; Williams et al., 2009; Bossers et al., 2010), we searched the miRanda and miRbase prediction algorithm databases (microRNA.org) for mRNA targets of these miRNAs that might mediate potentially compensatory pathways to promote neuronal viability. Surprisingly, all four miRNAs were predicted to target sirt1, a histone deacetylase involved in mediating neuronal cell stress responses (Brunet et al., 2004; Qin et al., 2006; Bonda et al., 2011). To determine whether target sirt1 transcripts were also differentially regulated in the frontal and temporal cortex in aMCI, we performed qPCR analysis of sirt1 expression levels in the same tissue samples used for miRNA analysis. This analysis revealed that sirt1 mRNA levels were significantly upregulated by ∼40% in the frontal cortex of aMCI compared to NCI and AD subjects (**Figure 1C**). By contrast, sirt1 levels were stable in temporal cortex across the diagnostic groups (**Table 4**). Furthermore, increased sirt1 mRNA expression was significantly associated with decreased miR-212 levels frontal cortex levels, but this association was not found in the temporal cortex (**Figure 2**).

### Localization of miR-23a and Sirt1 in the Frontal Cortex

Dual miR-23a in situ hybridization and sirt1 immunohistochemistry revealed nuclear miR-23a labeling in frontal cortex layer III neurons in tissue obtained from an 87 year-old female subject who died with a clinical diagnosis of aMCI (**Figure 3**). Sirt1 protein immunoreactivity was found primarily in the nucleus but also within the cytoplasm of the same neurons, providing evidence that miR-23a has direct access to its sirt1 target and that miR-directed sirt1 regulation in cortex is neuronal in origin.

### miR-212 and miR-23a Down-regulation Increases Sirt1 Protein Expression in Human Neuronal Cells

To determine the extent to which concomitant down-regulation of miR-212/132 and miR-23a/b and up-regulation of sirt1 observed in aMCI frontal cortex represented a functionally significant relationship, we treated human hNT neuronotypic cells with specific inhibitors of these miRNAs and measured sirt1 protein expression. Interestingly, inhibition of miR-212, miR-132, miR-23a, or miR-23b individually had no effect on sirt1 expression, but we found that concurrent inhibition of miR-212 and miR-23a resulted in a significant ∼100% increase in sirt1 (**Figure 4**), whereas co-inhibition of miR-132 and miR-23a resulted in a ∼40% increase in sirt1 (p < 0.05, data not shown). By contrast, miR-23b co-inhibition with miR-212 or miR-132 had no effect on sirt1 expression (data not shown).

quantitative analysis. \*\*p < 0.01 via One-way ANOVA with Bonferroni correction for multiple comparisons.

TABLE 4 | qPCR analysis of select miRNAs and sirt1 mRNA in temporal cortex of NCI, aMCI, and AD subjects.

\*p < 0.05, \*\*p < 0.01.

### miR-23a and miR-212 Down-regulation Protects against Aβ1−42 Induced Cell Death via Sirt1

To test whether miR-212 and miR-23a regulation of sirt1 results in neuronal protection, hNT cells were treated with inhibitors of either miRNA independently, or with inhibitors of both miRNAs combined, followed by challenge with Aβ1−42, which has been shown to induce cell death in these neuronal cells (Counts and Mufson, 2010). Inhibition of both miRNAs was sufficient to reduce Aβ1−42 induced cell death (**Figure 5**). This response was blocked by co-administration with the sirt1 inhibitor EX527 (**Figure 5**), supporting the concept that miRNA regulation of sirt1 expression is a viable neuroprotective pathway.

### DISCUSSION

The discovery of miRNAs (Lagos-Quintana et al., 2001) introduced a new layer of complexity to gene regulation, but also afforded the opportunity to better understand the molecular underpinnings of cellular function and dysfunction. This concept has already been well-demonstrated in cancer research and the potential therapeutic value of targeting miRNAs has been gaining acceptance in that field (Ma and Weinberg, 2008). With respect to neurodegenerative disorders such as AD, the presence of miRNAs in the human brain and evidence for miRNA function in a wide variety of complex neuronal processes, including synaptic plasticity, suggests that miRNA regulation could have immense implications not only for disease pathogenesis but also for neuronal responses to progressive neurodegeneration (Kosik, 2006). Prior studies show that miRNAs could impact AD progression through several mechanisms. For instance, postmortem human tissue studies in well-characterized cohorts have shown decreased neocortical levels of miR-29a/b, miR-9, and miR-107 in AD compared to control subjects, which was associated with increased BACE1 mRNA expression and Aβ generation (Hébert et al., 2008; Wang et al., 2008; Che et al., 2014); in particular, miR-107 is downregulated very early in the disease process (Wang et al., 2008). By contrast, miR-15a is decreased in AD brain compared to healthy controls and is implicated in tau hyperphosphorylation in vivo (Hébert et al., 2010). Here, we show that mir-132/212 and miR-23a/b are selectively down-regulated in the frontal cortex

in subjects clinically diagnosed with aMCI and that these alterations appear to be functionally linked to an up-regulation of sirt-1 and sirt-1 mediated protective responses. This novel finding adds to a growing literature on miRNA involvement in AD pathophysiology. However, rather than implicating another group of miRNAs in promoting neurodegeneration, our data support the concept that innate neuronal compensatory miRNA-mediated pathways are also activated in aMCI. A greater understanding of these and other miRNA pathways functioning during these putative prodromal stages of AD holds the promise that these pathways could be harnessed pharmacologically for drug development.

The miR-132/212 cluster has been implicated in several neuronal pathways, including dendritic elaboration (Magill et al., 2010) and learning and memory (Wang et al., 2013), and is downregulated in AD neocortex (Cogswell et al., 2008; Hébert et al., 2013; Lau et al., 2013; Wong et al., 2013). In particular, Wong and colleagues have previously shown that the miR-212/132 cluster is down-regulated in temporal cortex in AD, and that inhibition of miR-212 and/or miR-132 expression can induce apoptosis in primary neurons after 1 week in culture via activation of a foxo3a-mediated cell death pathway (Wong et al., 2013). In this regard, we replicated the finding that the miR-212/132 cluster is down-regulated in the temporal cortex in AD. However, we found no reductions in miR-212/132 in this region in aMCI; miR-212/132 was down-regulated in aMCI only in frontal cortex. In addition, in our hands experimental inhibition of miR-212 and/or miR-132 had no effect on cell survival of hNT neurons after 48 h in the absence of Aβ1−42. However, co-inhibition of either miR-212 or miR-132 with miR-23a conferred neuroprotection against Aβ1−42 in a sirt1 dependent manner. These discrepancies are likely explained by several factors. For instance, down-regulation of these miRNAs may have different functional implications in the AD temporal cortex, which displays a greater degree of degenerative changes than the aMCI frontal cortex. Notably, we found that sirt1 upregulation in frontal cortex was confined to aMCI, suggesting that other AD-related pathways prevent sirt1 activation as the disease progresses. The down-regulation of miR-212/132 in aMCI frontal cortex may initially participate in protective compensatory mechanisms, but with sustained reductions these miRNAs may join a pathological cascade that promotes disease progression through foxo3a. In this regard, sirt1 activity has been shown to protect against foxo3a-mediated pro-apoptotic pathways by the deacetylation of this transcription factor (Brunet et al., 2004; Qin et al., 2008). Moreover, the differential roles for this cluster may depend on target binding partners, since miR-212/132 and miR-23a co-inhibition was required for sirt1 activation and neuroprotection. Hence, the activity of the miR-212/132 cluster may be context dependent during the progression of AD.

miR-23a/b has also been shown to be dysregulated in the AD brain (Cogswell et al., 2008; Lau et al., 2013), yet much less is known about this miRNA cluster in neuronal function. However, like miR-212/132, miR-23a levels have been inversely linked to apoptosis. In a recent report detailing mechanisms for neuronal cell death in a model of traumatic brain injury, in vitro studies revealed that miR-23a inhibition

increased etoposide-induced cell death after 24 h in cortical neurons via caspase activation (Sabirzhanov et al., 2014). By contrast, we found that miR-23a inhibition alone had no effect on Aβ-induced cell death after 48 h, yet was neuroprotective in the presence of miR-212 by activating sirt1. Again, these discordances are not incompatible, but suggest that miR-23a has both pro-apoptotic and neuroprotective properties that depend on specific miRNA binding partners and whether these miRNAs are targeting pro-apoptotic factors such as caspases or, as presently shown, factors such as sirt1 that promote neuronal viability.

While the functional consequences of sirt1 up-regulation in this paradigm are unclear, sirt1 deacetylase activity plays an important role in regulating diverse cellular processes including aging, inflammation, and stress resistance (Imai et al., 2000; Brunet et al., 2004; North and Verdin, 2004; Herskovits and Guarente, 2014). In the adult brain, sirt1 can also modulate dendritic (Codocedo et al., 2012) and synaptic plasticity as well as memory formation (Gao et al., 2010; Michán et al., 2010). In addition to its importance during normal brain aging, sirt1 may also confer protective properties in neurodegenerative disorders such as AD (Qin et al., 2006, 2008; Min et al., 2010). Experimentally, sirt1 can reduce both Aβ-induced toxicity in neuronal cell lines (Conte et al., 2003; Han et al., 2004) and amyloid plaque formation in AD transgenic mice (Karuppagounder et al., 2009; Vingtdeux et al., 2010). Neuronal sirt1 expression has also been linked to non-amyloidogenic APP processing (Qin et al., 2006). Hence, we validated these concepts by showing that miR212/132 and miR-23a-mediated neuroprotection against Aβ is prevented by a sirt1-specific inhibitor. Sirt1 has also been shown to ameliorate tangle-like pathology in tau mutant mice (Kim et al., 2007; Min et al., 2010), possibly by tau deacetylation that allows ubiquitin ligases to target tau for degradation (Min et al., 2010; Cohen et al., 2011). Hence, there are several mechanisms by which miR-212/132 and miR-23a co-regulation of sirt1 expression may promote neuroprotection in the frontal cortex in aMCI. However, it is

FIGURE 5 | Experimental down-regulation of miR-212 and miR-23a protects against Aβ1−42 in a sirt1-dependent manner. Human hNT neuronotypic cells were co-transfected with small inhibitors of miR-212 and 23a or scrambled negative controls (CTL). Cells were then treated with 10µM Aβ1−42 for 48 h in the presence or absence of 100 nM EX527, a sirt1-specific inhibitor, followed by incubation with propidium iodide (PI) as an assay for cell death. Bar graph shows that Aβ1−42 treatment resulted in significant cell death that was significantly reduced by miR-212/23a co-repression. This rescue effect was reversed by EX527, suggesting that the protective effects of miR-212/23a co-repression was mediated by sirt1. n = 8/group; \*p < 0.05 vs. CTL; \*\*p < 0.05 vs. Aβ1−42 via One-way ANOVA with Bonferroni correction for multiple comparisons.

sirt1's role in the regulation and maintenance of dendritic growth (Codocedo et al., 2012) that could also play an important role in frontal cortex plasticity responses in MCI. In this regard, sirt1 knockout (KO) mice display reduced dendritic profiles relative to wild-type mice (Michán et al., 2010). Furthermore, overexpression of sirt1 can increase dendritic arborization, a phenotype that renders neurons resistant to dendritic dystrophy induced by Aβ (Codocedo et al., 2012). These data, taken together with multiple lines of evidence for a paradoxical up-regulation of synaptic elements within the frontal cortex during MCI (Counts et al., 2006; Bell et al., 2007; Williams et al., 2009; Bossers et al., 2010), suggest that sirt1 may mediate a synaptic remodeling response to mounting pathology during the progression of AD.

In summary, our data suggest that the transition from normal cognitive function in aging to a clinical diagnosis of aMCI may involve the suppression of brain microRNA networks such as miR212/132 and miR-23a. In the frontal cortex, this may result in the induction of cellular survival pathways including the stabilization of sirt1. Ultimately, as aMCI progresses to AD, the robust sirt1 expression is lost in frontal cortex, mimicking the situation seen in the temporal cortex. Recent microarray data suggest that the pattern of expression seen in sirt1 is also seen in other genes, particularly those involved in metabolic and synaptic machinery (Berchtold et al., 2013; Counts et al., 2013). Future research is necessary to confirm these findings in other AD cohorts, clarify whether the cumulative effect of these expression patterns is protective in vivo and by which sirt1-mediated mechanism, and determine whether augmentation of cell survival pathways such as sirt1 represents a viable strategy to delay neurodegeneration in prodromal AD. Linking these concepts to recent advances in miRNA knockdown and mimetic technology will usher in a new class of powerful in vitro and in vivo approaches to test the efficacy of miRNA-based therapies in preclinical AD models.

#### AUTHOR CONTRIBUTIONS

RW participated in the experimentation and manuscript preparation. EM participated in the case selection, data analysis, and manuscript preparation. SC participated in the

#### REFERENCES

study design, experimentation, data analysis, and manuscript preparation.

#### ACKNOWLEDGMENTS

The authors would like to thank the participants of the Rush Religious Orders Study for their continued altruism and support of AD research. This work was supported by NIH AG042146, NIH AG014449, NIH AG043375, The Barrow Neurological Foundation and Barrow Beyond, and the Saint Mary's Foundation of Mercy Health Saint Mary's Hospital, Grand Rapids, MI.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Weinberg, Mufson and Counts. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# EFhd2, a Protein Linked to Alzheimer's Disease and Other Neurological Disorders

Irving E. Vega\*

Department of Translational Science and Molecular Medicine, College of Human Medicine, Michigan State University, Grand Rapids, MI, USA

EFhd2 is a conserved calcium binding protein linked to different neurological disorders and types of cancer. Although, EFhd2 is more abundant in neurons, it is also found in other cell types. The physiological function of this novel protein is still unclear, but it has been shown in vitro to play a role in calcium signaling, apoptosis, actin cytoskeleton, and regulation of synapse formation. Recently, EFhd2 was shown to promote cell motility by modulating the activity of Rac1, Cdc42, and RhoA. Although, EFhd2's role in promoting cell invasion and metastasis is of great interest in cancer biology, this review focusses on the evidence that links EFhd2 to Alzheimer's disease (AD) and other neurological disorders. Altered expression of EFhd2 has been documented in AD, Parkinson's disease, Huntington's disease, Amyotrophic Lateral Sclerosis, and schizophrenia, indicating that Efhd2 gene expression is regulated in response to neuropathological processes. However, the specific role that EFhd2 plays in the pathophysiology of neurological disorders is still poorly understood. Recent studies demonstrated that EFhd2 has structural characteristics similar to amyloid proteins found in neurological disorders. Moreover, EFhd2 co-aggregates and interacts with known neuropathological proteins, such as tau, C9orf72, and Lrrk2. These results suggest that EFhd2 may play an important role in the pathophysiology of neurodegenerative diseases. Therefore, the understanding of EFhd2's role in health and disease could lead to decipher molecular mechanisms that become activated in response to neuronal stress and degeneration.

#### Edited by:

Tibor Hortobágyi, University of Debrecen, Hungary

#### Reviewed by:

Caterina Scuderi, Sapienza University of Rome, Italy Carlos Barcia, Universitat Autònoma de Barcelona, Spain Michael James Hurley, Imperial College London, UK

> \*Correspondence: Irving E. Vega<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Neurodegeneration, a section of the journal Frontiers in Neuroscience

Received: 19 January 2016 Accepted: 21 March 2016 Published: 31 March 2016

#### Citation:

Vega IE (2016) EFhd2, a Protein Linked to Alzheimer's Disease and Other Neurological Disorders. Front. Neurosci. 10:150. doi: 10.3389/fnins.2016.00150 Keywords: EFhd2, tau, neurodegenerative disease, neurological disorders, Alzheimer's disease

## INTRODUCTION

Efhd2 gene codes for a 26.8 kDa highly conserved protein, from nematodes to human, located in chromosome 4 (4E1;4 74.75 cM) in mice and chromosome 1 (1p36.21) in humans. EFhd2 was first identified in a proteomics screen designed to discriminate CD8 from CD4 and CD19 lymphocytes. The abundance of this novel protein was found to be reduced in CD4 and C19 in comparison to CD8 lymphocytes (Vuadens et al., 2004; Dütting et al., 2011). In this original study, the novel protein was named Swiprosin 1, in reference to the Swiss-Prot database used for the tandem mass spectrometry data analysis (Vuadens et al., 2004). Subsequently, the name of this novel protein was changed to EF-hand domain family, member D2 (EFhd2) due to the presence of two EF-hand calcium binding motifs. Characterization of EFhd2 demonstrated that it is an ubiquitous calcium binding protein, preferentially expressed in the central nervous system (Avramidou et al., 2007; Vega et al., 2008; Hagen et al., 2012; Ferrer-Acosta et al., 2013b). Further sequence analyses indicated that EFhd2 has a coiled-coil domain at the C-terminus, which is a conserved domain among fibrillar proteins and required for protein–protein interaction (Ferrer-Acosta et al., 2013a). At the N-terminus, EFhd2 has a distinctive polyalanine motif that varies in size (between 6 and 9 alanines) and it is only present in mammals (Dütting et al., 2011; Ferrer-Acosta et al., 2013a). The function of EFhd2's polyalanine motif is still unknown, but proteins containing polyalanine expansions have been shown to be linked to different neurological disorders (Albrecht and Mundlos, 2005). However, the physiological and pathological roles of EFhd2 are still poorly understood.

EFhd2 may function as a signaling or cytoskeleton regulatory protein. In WEHI231 cells, it was shown that EFhd2 is required for the regulation of the canonical NFkB pathway upon activation of the B-cell receptor (BCR; Avramidou et al., 2007; Hagen et al., 2012; Kim et al., 2013). ShRNA-mediated EFhd2 knockdown led to increase IkB phosphorylation, which is a prerequisite for translocation of NFkB to the nucleus, upon BCR activation (Avramidou et al., 2007; Kim et al., 2013). Based on this result, the authors suggested that EFhd2 may play a role as negative regulator of NFkB in the BCR signaling pathway (Avramidou et al., 2007). Alternatively, another study found EFhd2 at the plasma membrane, where it facilitates the assembly of the BCR and appears to work as a scaffold protein required for the function of Syk, SLP-65, and PLCγ2 during BCR-induced calcium flux (Kroczek et al., 2010). The functional interaction of EFhd2 with BCR signaling pathway and modulation of IkB phosphorylation indicates a potential regulatory role in cell survival or fate. Interestingly, EFhd2 was identified as a novel pro-caspase-9-interacting protein in H460 cells (Ch˛ecinska ´ et al., 2009). EFhd2 association with (inactive) pro-caspase-9 protein suggests that it may regulate the activation of apoptosis (Ch˛ecinska et al., 2009 ´ ). In contrast, other reports indicated that EFhd2 mediates actin bundling and regulates cell spreading and migration (Huh et al., 2013; Kwon et al., 2013). Moreover, recent studies indicated that EFhd2 was upregulated by epidermal growth factor signaling and mediates cell migration through the modulation of Rac1, Cdc42, and RhoA activity (Huh et al., 2015). Although, the direct role of EFhd2 in regulating cellular survival and apoptosis is still unclear, these results indicate that this novel protein plays an important role in modulating cellular responses elicited by different environmental cues, such as trophic factors or stimulating antibodies (Avramidou et al., 2007; Huh et al., 2015).

EFhd2 has also been associated with different pathological processes, from cancer to neurological disorders (**Table 1**). In cancer, EFhd2 was found to be overexpressed in the majority of carcinomas, colon cancer, and melanoma (Huh et al., 2015). The results indicated that EFhd2 may mediate invasion and metastasis of cancerous cells, suggesting it plays an important role in the biology of cancer. EFhd2 gene expression and protein abundance have been also shown to be altered in AD and other neurological disorders. Thus, even though the role that EFhd2 plays in cancer biology is of great interest, this review focusses on EFhd2's role in the central nervous system, specifically its association with AD and other neurological disorders.

### IS EFHD2 A MODULATOR OF FUNCTIONAL SYNAPSE FORMATION?

EFhd2 is highly expressed in neurons compared to other cell types of the central nervous system, where may play an important role in synapse formation (Reynolds et al., 2008; Vega et al., 2008; Ferrer-Acosta et al., 2013b; Borger et al., 2014). EFhd2 proteins were found in the cytosol and proximal to the membrane in neurons of most brain regions, including higher expression in the deeper layers of the cortex and hippocampus (Ferrer-Acosta et al., 2013b; Borger et al., 2014). EFhd2 co-localized with neurite markers such as tau, MAP2,synapsin, and PSD95, suggesting that its neuronal function could be associated with vesicle transport and synapse homeostasis (Borger et al., 2014; Purohit et al., 2014). Consistent with this putative function, a recent study showed in vitro that knockdown of EFhd2 increased synpasin 1a/b puncta labeling in neurites, suggesting that modulation of EFhd2 affects the development of functional synapses, but it had no effect on converting them to mature synapses as determined by the colocalization of synapsin and PSD95 (Borger et al., 2014). These results imply that EFhd2 may modulate the formation of new synapses, a process that it is relevant for different brain functions, such as learning and memory.

Characterization of an Efhd2 knockout mouse provided further insights about its function in the central nervous system. Purohit et al. (2014) showed that deletion of Efhd2 gene has no detectable effect on brain anatomy or function. Interestingly, they showed that vesicle transport velocity was enhanced in Efhd2(−/−) knockout primary hippocampal neurons and that EFhd2 protein inhibited kinesin mediated microtubule gliding in vitro (Purohit et al., 2014). The authors proposed that EFhd2 may interfere with the interaction between kinesin and microtubules (Purohit et al., 2014). In this regard, EFhd2 has been showed to co-purify with tubulin in the synaptosome fraction as well as to mediate actin bundling (Huh et al., 2013; Kwon et al., 2013; Purohit et al., 2014). Although, there is no evidence that Efhd2 gene knockout affects brain function, these results suggest that EFhd2 may serve as a modulator of synapse formation by regulating the velocity of vesicle transport and cytoskeleton rearrangement.

### EFHD2 IN ALZHEIMER'S DISEASE

EFhd2 protein abundance was found altered in AD, suggesting that Efhd2 gene expression may be regulated in response to neurodegeneration (Vega et al., 2008; Borger et al., 2014; **Table 1**). Two different studies showed that the chromosome region encompassing the Efhd2 gene locus is linked to (and a third study showed that it is associated with) late-onset Alzheimer's disease (LOAD; Hiltunen et al., 2001; Myers et al., 2002; Holmans et al., 2005). The linkage between Efhd2 gene and LOAD is yet to be determined; however, we and others have shown that EFhd2 expression and protein abundance is altered in AD and

#### TABLE 1 | Identification of EFhd2 associated with neurological disorders.

animal models that mimic the pathophysiology associated with AD (Vega et al., 2008; Tseveleki et al., 2010; Borger et al., 2014). Previously, we showed that EFhd2 protein is increased in the tauopathy mouse model JNPL3, which expresses the human P301L mutant tau protein (Vega et al., 2008; Ferrer-Acosta et al., 2013b). We showed that EFhd2 protein abundance increased as the accumulation of pathological tau and progression of the motor impairment phenotype augmented in JNPL3 mice (Vega et al., 2008). In an independent expression profiling study, using a different AD mouse model, APP23, EFhd2 was also found to be overexpressed (Tseveleki et al., 2010). It is important to mention that the APP23 AD mouse model overexpresses human Amyloid Precursor Protein with double Swedish mutation (K670M/N671L) and it has been shown that tau mediates the toxicity observed in this mouse model (Sturchler-Pierrat et al., 1997; Ittner et al., 2010). These results suggest that EFhd2 is upregulated in response to pathological processes associated with tau-mediated neurodegeneration.

EFhd2 protein increased abundance in mouse models of neurodegeneration was validated in postmortem AD brain (Vega et al., 2008; Ferrer-Acosta et al., 2013b). Using quantitative immunoblotting, we demonstrated that EFhd2 protein abundance is increased in postmortem frontal cortices in AD cases (Ferrer-Acosta et al., 2013b). In contrast, Borger et al. (2014) showed that EFhd2 protein abundance is reduced in AD cases and other dementias, including frontotemporal lobar degeneration with TDP43 pathology. Moreover, they showed that EFhd2 mRNA was also significantly lower in AD, when compare to normal aging controls. The discrepancy between these two reports could be due to differences in postmortem brain sample selection, agonal stage, activated glial cells, and/or differential immune cell infiltration. For example, in our study, we included only AD postmortem brain with Braak stage IV or higher (Ferrer-Acosta et al., 2013b). In Borger et al. (2014), Braak stage is reported for only a few of the cases used. It is important to mention that in those reported cases with Braak stage VI, EFhd2 protein level is higher than in the other AD cases that the Braak stage was not indicated. Other technical reasons, such as anti-EFhd2 antibodies used, protein extraction method and differential subcellular localization, could contribute to the discrepancy. Interestingly, Twine et al. (2011) identified EFhd2 as one of the genes with increased alternative splicing in frontal cortices of AD patients. This is consistent with previous results that identified two protein bands corresponding to EFhd2, which might contribute to the detection of different EFhd2 isoforms depending on the antibody used (Avramidou et al., 2007; Vega et al., 2008). Nevertheless, what is consistent among these studies is that EFhd2 expression was found altered in AD. Further studies are required to determine the molecular mechanisms involved in the regulation of the Efhd2 gene and protein under pathological or cellular stress conditions.

Further biochemical characterization of EFhd2 protein demonstrated that this novel protein has the molecular and structural features of amyloid proteins (Ferrer-Acosta et al., 2013b). In vitro studies showed that Thioflavin S (a dye that selectively binds to amyloid structures) binds recombinant EFhd2 protein, indicating that EFhd2 transition from a mostly helical and random coil structure to cross-beta-sheet (Ferrer-Acosta et al., 2013a,b). Amyloid proteins tend to form oligomers and filamentous structures. Electron microscopy analyses confirmed that EFhd2 forms filaments in vitro without the requirement of a nucleation factor (Congdon et al., 2008; Ferrer-Acosta et al., 2013b). Furthermore, the presence of calcium reduces EFhd2's ability to form filaments, and the coiledcoil domain was shown to be required for the formation of EFhd2 homodimers (Ferrer-Acosta et al., 2013b). These results indicated that formation of EFhd2 filaments could promote the association with pathological tau filaments. To test this hypothesis, immunohistological analyses of AD brain slices were performed. The results showed that EFhd2 co-localized with PHF1 (an antibody that recognized tau filaments) in the somatodendritic compartment, validating the association of EFhd2 with filamentous tau structures (Ferrer-Acosta et al., 2013b). Moreover, immune-gold electron microscopy showed that EFhd2 and tau, purified from AD brain, formed co-filaments (Ferrer-Acosta et al., 2013b). Moreover, in vitro protein–protein interaction assays demonstrated that EFhd2's coiled-coil domain is necessary for its association with tau proteins purified from brain extract derived from JNPL3 mice (Ferrer-Acosta et al., 2013b). Thus, it is plausible to hypothesize that formation of EFhd2 oligomers may serve as nucleation factor for tau oligomerization and, consequently, NFTs in AD and other tauopathies (**Figure 1**). Nevertheless, further studies are required to determine EFhd2's capability to enhance protein aggregation in tauopathy and other neurodegenerative disorders.

## EFHD2 IS ASSOCIATED WITH PARKINSON'S DISEASE AND OTHER NEUROLOGICAL DISORDERS

EFhd2 has been also found associated with Parkinson's disease (PD). Liscovitch and French (2014) showed that EFhd2 expression is increased in substantia nigra in PD. In this study, they found that EFhd2 overexpression correlated with the positive co-expression of α-synuclein and IFNγ, establishing a molecular relationship between PD and inflammation (Liscovitch and French, 2014; **Table 1**). In contrast, EFhd2 protein abundance was found reduced in the substantia nigra of a mouse model where PTEN-induced kinase 1 (PINK1) has been knockout (Diedrich et al., 2011). PINK1 is a mitochondrial protein, the loss-of-function mutation of which induces early-onset PD. These conflicting reports could be due to the difference between mechanisms associated with sporadic PD cases and those underlying the phenotype of the PINK1 knockout mice. However, in vitro studies identified EFhd2 as a protein secreted/released by microglia cells upon incubation with nitrated α-synuclein (Reynolds et al., 2008). This is the first study that shows expression of EFhd2 in microglia cells, but, more importantly, it also indicates that EFhd2, a cytosolic protein, could be released from cells in response to a pathological signal. Taken together, EFhd2 altered protein abundance in PD provides strong evidence that it is involved in molecular mechanisms associated with neurodegeneration.

EFhd2 associates with known proteins linked to the pathophysiology of PD (**Table 1**). In a proteomics approach, EFhd2 was identified associated with leucine-rich repeat kinase 2 (Lrrk2), the most common causative gene of inherited PD (Meixner et al., 2011). Interestingly, this study demonstrated that Lrrk2 decreased actin polymerization, promoting the accumulation of monomeric actin (G-actin; Meixner et al., 2011). Knockdown of Lrrk2 affects actin cytoskeleton and cellular morphology, indicating that Lrrk2 plays a direct role on actin (Meixner et al., 2011). In contrast, EFhd2 promotes actin bundling (Huh et al., 2013). Thus, these results suggest that Lrrk2 and EFhd2 may compete for the interaction with F-actin, as regulators of actin cytoskeleton dynamics. Moreover, Lrrk2 has been shown to phosphorylate tau and promote tau aggregation, as observed in postmortem brain tissue from PD cases with Lrrk2 mutations (Guerreiro et al., 2015). On the basis of their link to neurodegeneration, it is reasonable to speculate that there could be a pathological connection between Lrrk2 and tau through EFhd2.

EFhd2 altered protein abundance and its association with pathological proteins have also been found in other neurological disorders. EFhd2 protein was identified as a co-aggregating

protein with poly-GA C9orf72, a common pathogenic protein in amyotrophic lateral sclerosis (ALS) and frontotemporal lobar degeneration (FTLD; May et al., 2014). This result suggests that EFhd2 may be involved in the molecular mechanisms that lead to aggregation of pathological proteins, other than tau, in proteinopathies such as ALS. Consistently, EFhd2 was one of the uniquely identified proteins found in lipid rafts isolated from a mouse model that expresses the G93A SOD1 mutant; a mutation found in inherited ALS cases (Zhai et al., 2009). This result indicates that EFhd2 subcellular localization could also be altered in neurodegeneration.

In addition to its association with neurodegenerative diseases, Efhd2 gene expression has also been found altered in psychiatric disorders. Two independent studies demonstrated that EFhd2 is up-regulated in schizophrenia (Martins-de-Souza et al., 2009, 2010). Postmortem analysis of dorsolateral prefrontal cortex and mediodorsal thalamus, two brain regions associated with the pathophysiology of schizophrenia, revealed a significant increase in EFhd2 protein level in comparison to normal control cases (Martins-de-Souza et al., 2009, 2010). In contrast, EFhd2 was found down-regulated in prefrontal cortex and up-regulated in amygdala of individuals that committed suicide (Kékesi et al., 2012). These results suggest that altered levels of EFhd2 are directly associated with brain pathology.

#### CONCLUSION

In summary, EFhd2 harbors similarities to known and wellstudied neuropathological proteins. For instance, EFhd2 is a structural disorder protein mainly composed of random coils and alpha helices, forms filamentous structures, associates with vesicle trafficking and cytoskeleton rearrangement, co-aggregate, and co-purify with pathological proteins, its expression and protein abundance is altered in neurodegenerative diseases. Based on these similarities and the published results discussed

above, it is plausible to hypothesize that neuronal stress due to environmental cues (such as cytokines) or pathological protein aggregation induces changes in Efhd2 gene expression (**Figure 1**). In this context, activation of signaling proteins, such as Cdk5 (Vázquez-Rosa et al., 2014), induces posttranslational modifications on EFhd2 that affect its calcium binding activity and promotes self-oligomerization (**Figure 1**). The accumulation of EFhd2 oligomers could serve as a nucleation factor of tau proteins facilitating its accumulation at the somatodendritic compartment, which also affects kinesin-mediated fast axonal transport (**Figure 1**; Kanaan et al., 2013). Consequently, the accumulation of EFhd2 proteins and tau aggregates would lead to activation of apoptosis (**Figure 1**). Interestingly, up-regulation of EFhd2 could also promote aberrant actin bundling, leading to the formation of Hirano bodies in AD (**Figure 1**, Sonoda et al., 2015).

Alternatively, EFhd2 altered expression in neurological disorders could be related to neuroprotection. The identification of EFhd2 co-localization with neurofibrillary tangles (NFTs) in the somatodendric compartment and its filament formation capability suggest that EFhd2 may affect the kinetic of tau filament formation, promoting the generation of stable ultrastructures (**Figure 1**). The generation of NFTs may sequester tau oligomers, preventing the spread of toxic tau to the interstitial fluid. Concomitantly, EFhd2's accumulation in NFTs induces a loss-of-function effect that promotes the generation of new functional synapses

#### REFERENCES

through increase vesicle transport velocity similar to that observed in Efhd2 knockout mice. This may explain why NFT bearing neurons are still synaptically active and functionally integrated in neuronal circuits (Kuchibhotla et al., 2014). Nevertheless, further studies are crucial to determine the role that EFhd2 plays in the pathophysiology of neurological disorders.

#### AUTHOR CONTRIBUTIONS

IEV revised the literature and wrote the manuscript.

#### FUNDING

The work reviewed was supported, in part, by funding from National Institute of Neurological Disorders and Stroke (1SC1NS066988, 8R25NS080687).

#### ACKNOWLEDGMENTS

The author would like to acknowledge the contribution of graduate and undergraduate students in the quest to characterize the pathological and physiological role of EFhd2. EFhd2 characterization was supported, in part, by NIH-NINDS grant 1SC1NS066988 to IEV. The students were supported by NIH-NIGMS 5R25GM061151 and NIH-NINDS 8R25NS080687 training grants.

with pathological tau in Alzheimer's disease. J. Neurochem. 125, 921–931. doi: 10.1111/jnc.12155

abnormalities meet the axonal transport system. Exp. Neurol. 246, 44–53. doi: 10.1016/j.expneurol.2012.06.003

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Vega. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# A Rapid, Semi-Quantitative Assay to Screen for Modulators of Alpha-Synuclein Oligomerization Ex vivo

Marion Delenclos 1 , Teodora Trendafilova1 , Daryl R. Jones 1 , Simon Moussaud1 , Ann-Marie Baine1 , Mei Yue1 , Warren D. Hirst 2 and Pamela J. McLean1, 3 \*

*1 Department of Neuroscience, Mayo Clinic, Jacksonville, FL, USA, 2 Pfizer Neuroscience Research Unit, Cambridge, MA, USA, 3 Mayo Graduate School, Mayo Clinic, Jacksonville, FL, USA*

Alpha synuclein (αsyn) aggregates are associated with the pathogenesis of Parkinson's disease and others related disorders. Although modulation of αsyn aggregation is an attractive therapeutic target, new powerful methodologies are desperately needed to facilitate *in vivo* screening of novel therapeutics. Here, we describe an *in vivo* rodent model with the unique ability to rapidly track αsyn-αsyn interactions and thus oligomerization using a bioluminescent protein complementation strategy that monitors spatial and temporal αsyn oligomerization *ex vivo*. We find that αsyn forms oligomers *in vivo* as early as 1 week after stereotactic AAV injection into rat substantia nigra. Strikingly, although abundant αsyn expression is also detected in *striatum* at 1 week, no αsyn oligomers are detected at this time point. By 4 weeks, oligomerization of αsyn is detected in both striatum and substantia nigra homogenates. Moreover, in a proof-of-principle experiment, the effect of a previously described Hsp90 inhibitor known to prevent αsyn oligomer formation, demonstrates the utility of this rapid and sensitive animal model to monitor αsyn oligomerization status in the rat brain.

#### Edited by:

*Irving E. Vega, Michigan State University, USA*

#### Reviewed by:

*Alessandro Bertoli, University of Padova, Italy Kathleen A. Maguire-Zeiss, Georgetown University Medical Center, USA*

> \*Correspondence: *Pamela J. McLean<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Neurodegeneration, a section of the journal Frontiers in Neuroscience*

Received: *04 September 2015* Accepted: *21 December 2015* Published: *19 January 2016*

#### Citation:

*Delenclos M, Trendafilova T, Jones DR, Moussaud S, Baine A-M, Yue M, Hirst WD and McLean PJ (2016) A Rapid, Semi-Quantitative Assay to Screen for Modulators of Alpha-Synuclein Oligomerization Ex vivo. Front. Neurosci. 9:511. doi: 10.3389/fnins.2015.00511* Keywords: alpha-synuclein, oligomers, protein-fragment complementation, bioluminescence, viral vector model

## INTRODUCTION

Under pathological conditions alpha-synuclein (αsyn) can misfold and aggregate into insoluble deposits that accumulate in cells to toxic levels. The conversion of αsyn from its functional conformation into a misfolded and toxic conformation constitutes the basis of a group of diseases known as synucleinopathies which include Parkinson's disease (PD), multiple system atrophy, and dementia with Lewy bodies (Goedert and Spillantini, 1998; Kim et al., 2014). Several lines of evidence demonstrate a strong association between αsyn aggregation and neurodegeneration (Irizarry et al., 1998; Chartier-Harlin et al., 2004; Winner et al., 2011). Fibrillar forms of αsyn are the major component of glial cytoplasmic inclusions, Lewy bodies (LBs), and Lewy neurites, defined as intracytoplasmatic inclusions and considered the pathological hallmarks of synucleopathies (Forno, 1996; Spillantini et al., 1997; Halliday et al., 2011).  The processes that lead to pathological aggregate formation occur through the formation of several soluble oligomeric intermediates that mature into the insoluble amyloid fibrils found in LBs. It has been proposed that prefibrillar forms of αsyn are the diseasecausative toxic species, while the insoluble fibrils might represent a protective pathway for surviving cells (Ross and Poirier, 2004; Paleologou et al., 2009; Winner et al., 2011; Kalia et al., 2013).

Although the mechanisms of αsyn-induced toxicity remain unclear, inhibition of oligomerization represents an attractive therapeutic approach. Such a strategy requires the availability of powerful cellular and rodent models. Current methods for detection of αsyn oligomers are challenging due to their dynamic nature and sensitivity to external conditions (Uversky, 2003; Drescher et al., 2012; Gurry et al., 2013) Thus, far, oligomer studies have used indirect methods and biochemical techniques that prohibit the study of αsyn oligomerization in a live cellular environment or in intact brain. More recently, protein fragment complementation assay (PCA) strategies that allow the detection of αsyn-αsyn interactions using fluorescence or bioluminescent reporters, have been widely developed and have been successfully applied to monitor αsyn oligomers not only in living cells (Outeiro et al., 2008; Putcha et al., 2010; Danzer et al., 2012) but also in rodent brain (Dimant et al., 2013, 2014; Aelvoet et al., 2014; McFarland et al., 2014).

Here, we generated and characterized a viral vector rodent model where humanized Gaussia princeps luciferase (hGluc) is used as a surrogate reporter of αsyn oligomerization in vivo in a fast, sensitive, and semi quantitative assay. We demonstrate that this rodent model can be utilized to track αsyn oligomerization in vivo and validate the potential use of the model by testing a novel Hsp90 inhibitor compound, known to reduce αsyn oligomerization in vitro, on ourin vivo bioluminescence read out.

## MATERIALS AND METHODS

### Viral Vector Production

The viral vectors pAAV-CBA-synuclein-LUC1-WPRE (SL1) and pAAV-CBA-SYNUCLEIN-luc2-WPRE (SL2) were constructed by inserting the human SNCA cDNA (h αsyn) fused to either the N-terminus half of humanized G. princeps luciferase (hGluc) (Tannous et al., 2005) or the C-terminus half of hGluc, into the EcoRV and NheI sites of the pAAV-CBA-WPRE. Viral vector pAAV-CBA-G. princeps luciferase was constructed by inserting the full length of hGluc gene into the XhoI and NheI sites of pAAV-CBA-WPRE vector. Adeno-associated virus (AAV) serotype 2/8 was produced by plasmid transfection with helper plasmids in HEK293T cells. Forty-eight hours later, the cells were harvested and lysed in the presence of 0.5% sodium deoxycholate and 50µ/ml Benzonase (Sigma-Aldrich, St. Louis, MO) by freeze-thawing, and the virus was isolated using a discontinuous iodixanol gradient. The genomic titer of each virus was determined by quantitative PCR.

#### Surgical Procedure

Adult Female Sprague Dawley rats (225–250 g, Harlan, USA) were housed and treated in accordance with the NIH Guide for Care and Use of Laboratory animals. All procedures were approved and conducted in accordance with the Mayo Clinic Institutional Animal Care and Use committee. Rats were housed 3 per cage with ad-libitum access to food and water during a 12 h light/dark cycle. Surgery was conducted under 2% isoflurane anesthesia mixed with O2 and N2 using a stereotaxic frame and a 10µl Hamilton syringe fitted with a 30 gauge needle. The scalp was exposed and a unilateral injection targeting the substantia nigra (SN) was performed at coordinates 5.2 mm posterior and 2 mm lateral to bregma, and 7.2 mm ventral relative to dura. AAV8 vectors were normalized by titer and volume, resulting in injection of an equal amount of genomes per copy per vector. Two microliter of a mixture of two viruses (SL1 8.1012 g/ml + SL2 8.1012 g/ml) were injected at a rate of 0.4µl/min using a microinjection pump (Stoelting Co, Wood Dale, IL). Control animals were injected with one virus only (2µl of AAV8-SL2 at 16.1012 genome/ml), or received one injection of 1µl of SL1 (8.1012 g/ml) in the SN of the left hemisphere (ML: −2 mm) and one injection of 1µl of SL2 (8.1012 g/ml) in the SN of the other hemisphere (ML: 2 mm). At the end of injection the needle remained in place for 5 min before being slowly retracted. Animals were then sutured with metal clips and monitored until fully recovered.

### Tissue Processing

For histological analyses, animals were deeply anaesthetized at 1 and 4 weeks post-injection with pentobarbital and perfused transcardially with ice-cold 0.9% saline, followed by 4% paraformaldehyde (PFA). Brains were removed and postfixed for 4 h in the same solution and were then transferred overnight to 25% sucrose solution for cryoprotection. The brains were cross-sectioned using a freezing stage sliding microtome (Leica SM2010R, Germany) at 40 µm in the coronal plane. Brains from a subset of animals from 1 to 4 weeks post injection were harvested fresh without fixation. The two hemispheres were separated and the striatum (STR) and midbrain from both sides were quickly dissected on ice and used directly for biochemical analysis.

### Immunohistochemistry

Immunohistochemical analysis was performed on free-floating sections (40µm) using primary antibodies against tyrosine hydroxylase (TH) (1:3000, MAB318, Millipore), and hαsyn (1:2000, 4B12, specific antibody to human form of αsyn, Covance). Sections were washed with phosphate buffer saline (PBS) before each incubation. After the initial wash, the sections were quenched for 20 min at RT with PBS solution, supplemented with 3% H2O2 (v/v) (Sigma-Aldrich), and 10% Methanol (v/v) (Pharmaco-Aaper) and were then blocked with 5% goat serum (Life technologies, Carlsbad, CA, USA) in T-PBS (0.25% Triton dissolved in PBS). The sections were incubated overnight at RT with one of the indicated antibodies, diluted in 2.5% goat serum and T-PBS. After washing, the brain slices were treated with 1:200 dilution of the appropriate biotinylated secondary antibody (Vector Laboratories, Burlingame, CA, USA) in 1% goat serum and T-PBS. Thereafter, the sections were rinsed and incubated with avidin-biotin-peroxidase complex (ABC peroxidase kit, Thermo Scientific). The staining was visualized by 3,3-diaminobenzidine (DAB; Sigma-Aldrich) used as a chromagen and activated by 1% H2O2. The sections were then mounted on gelatin-coated slides, dehydrated with alcohol, cleared with xylene and coverslipped with DePex mounting medium (Sigma-Aldrich).

### Stereology

Nigrostriatal cell loss was assessed using an unbiased stereology method using the optical fractionator principle as described previously (West et al., 1991). Lower magnification was used to trace the region of interest (SN in each hemisphere) at all levels in the rostro-caudal axis in 9–10 sections per animals. TH+ cells in the ventral tegmental area (VTA) were excluded. The counting was performed using the Stereo Investigator software (MBF bioscience). In addition, the counting frame size was adjusted so that ∼100–200 TH+ positive cells were counted in each side of the SN. The estimation of total neuron numbers was performed by using the optical fractionator formula and coefficient of error <0.1 was accepted (West, 1999).

## Densitometry

Images of forebrain were captured using the Aperio slide scanner (Aperio, Vista, CA, USA). The optical density of striatal TH+ fibers was measured using ImageJ software (Version 1.46, National Institutes of Health) at four coronal levels according to the rat brain atlas of Paxinos and Watson (6th edition, 2007): AP: +1.2,+ 0.8, 0.0 and −0.4 mm relative to bregma. The measured values were corrected for nonspecific background staining by subtracting values obtained from adjacent cortical areas. The data are expressed as a percentage of the corresponding area from the intact side.

### Ex vivo Luciferase Complementation Assay

Fresh STR and SN were homogenized briefly in 200µl cold PBS buffer using a Teflon pestle. One-hundred microliter of the homogenate was transferred to a 96 well plate (Costar, Corning, NY, USA) and the remainder was preserved for biochemical analysis. Luciferase activity from αsyn-αsyn interaction was measured in an automated plate reader (Victor 3 multilabel counter, Perkin Elmer, Waltham, MA, USA) at 480 nm following the injection of 100µl of the substrate coelenterazine (40µm, NanoLight tech, AZ, USA) with a signal integration of 2 s.

### Human αsyn Elisa

Quantitative analysis of hαsyn concentration in rat tissue was performed using hαsyn specific ELISA (#KHBOO61, Life tech) according to the manufacturer's instructions. Absorbance at 450 nm is directly proportional to the concentration of hαsyn present in the original specimen. αSyn concentrations were normalized to the total amount of protein in the homogenate. Protein content was measured using Bicinchoninic acid assay (BCA) (Thermo Fisher Scientific, Waltham, MA, USA).

### Western Blotting

Striatal and midbrain samples were suspended in RIPA buffer (Millipore, 20-188) containing protease inhibitor (Roche Diagnostics), homogenized on ice, and centrifuged at 13,000 × g for 10 min. Protein concentration of each lysate was determined by BCA. Twenty microgram proteins was loaded on a denaturing 4–12% Bis- Tris gradient gel (Nupage, Novex Gel, Life tech) and run according to the manufacturer's instructions. Subsequently, separated protein was transferred to polyvinylidene difluoride (PVDF) membrane, blocked in 5% non-fat milk in TBS-T, and immunoblotted for Hsp70 (1:5000, rabbit, ADI-SPA-812, Enzo). Anti-actin (1:10000, rabbit polyclonal, A2668, Sigma-Aldrich) was used as a loading control. All membranes were then incubated with a secondary antibody conjugated to HRP for 1 h at room temperature. Immunoreactivity was visualized using an ECL chemiluminescent detection Kit (Thermo Fisher Sci) and images were acquired with a CCD imaging system (LAS-4000, Fujifilm, Japan).

### SNX-9114 Treatment In vivo

SNX-9114 (Pfizer, PF-04944733) was described previously (McFarland et al., 2014) and was dissolved in a 0.5% methylcellulose solution before use. SNX-9114 was administered at a dose of 2 mg/kg by oral gavage twice for 1 week in a volume of 1 mL. As a control, the same volume of vehicle (0.5% methylcellulose) was simultaneously administered in a separate group of rats (vehicle group). SNX-9114 administration started 2 days after the stereotactic viral injection.

### Cell Cultures

H4 neuroglioma cells (ATCC, HTB-148, Manassas, USA) were maintained in Opti-MEM supplemented with 10% Fetal Bovine Serum (FBS). H4 cells were transfected with full length hGluc using Superfect transfection reagents (Qiagen, Chatsworth, CA, USA) with an equimolar ratio of plasmids according to the manufacturer's instructions.

A tetracycline-driven stable H4 cell line co-expressing hαsyn tagged with either the amino-terminal (SL1) or the carboxyterminal fragments (SL2) of hGluc luciferase was generated and described previously (Moussaud et al., 2015). SL1SL2 cells were cultured in Opti-MEM supplemented with 10% FBS, 200µg/ml geneticin, 200µg/ml hygromycin B, and 1µg/ml tetracycline to block the expression of the transgenes (SL1 and SL2). Cells were maintained at 37◦C in a 95% air/5% CO2 humidified incubator and all the cell cultures reagents were from Life technologies.

SL1SL2 stable cells were seeded into 6-well plates at the density of 1 × 105 cells/well and in a 96-well plate at the density of 3 × 104 cells/well in the absence of tetracycline. Twenty-four hours later, media was replaced with FBS-free media supplemented with SNX-9114 (25, 50, 100, or 200 nM) or with DMSO alone as control. After 24 h treatment, cells in the six well plates were lysed (150 mM NaCl, 5 mM Trisbase, 0.1% TritonX, cOmplete Mini, pH 7.4) and analyzed by western blotting using antibodies against: hαsyn (1:1000,mouse, 610787, Becton Dickinson, NJ, USA), Hsp70 (1:5000, rabbit, ADI-SPA-812, Enzo life), and actin (1:10000; rabbit polyclonal, A2668, Sigma-Aldrich), whereas cells in 96 well plates were used for luciferase activity measurement as described previously (Putcha et al., 2010).

#### Statistics

Data were analyzed using GraphPad Prism 6 (San Diego, CA) and are presented as mean ± standard error of the mean (S.E.M.). Statistical significance was determined using a Student's t-test or one-way analysis of variance with Tukey's multiple comparison post-hoc. P < 0.05 was considered significant.

### RESULTS

#### Ex vivo Monitoring of αsyn Oligomers using Bioluminescent PCA

The bioluminescent αsyn PCA strategy has been widely used by our lab and others (Outeiro et al., 2008; Putcha et al., 2010; Danzer et al., 2011; Moussaud et al., 2015) to investigate αsyn oligomerization in living cells. Briefly, two hαsyn proteins fused to N- or C-terminal halves of a reporter protein can reconstitute the enzymatic activity of the reporter when αsynαsyn interactions occur, thus providing a readout of αsynαsyn interactions and oligomerization. In this study, two AAV8 vectors expressing hαsyn fused with either the N-terminus or C-terminus of hGluc (referred hereafter as AAV-SL1 and AAV-SL2) were stereotactically co-injected unilaterally into the SN of adult rats to establish the in vivo bioluminescent PCA. Previous studies from our lab have characterized the oligomeric species formed in the bioluminescent complementation assay as a heterogeneous population of soluble dimers and higher order multimers (Danzer et al., 2011; Dimant et al., 2013). The term oligomer is used henceforth to represent the heterogenous population.

Delivery of AAV-SL1 and AAV-SL2 into the SN of the rat produces widespread overexpression of hαsyn along the nigrostriatal pathway as revealed by histological analysis (**Figures 1A,B**). Immunostaining with an antibody specific for hαsyn shows numerous cell bodies in the SN expressing the transgenes at 1 week and at 4 weeks while the non-injected side is devoid of hαsyn and is used as an internal control (**Figure 1A**). Transport of virally transduced αsyn is indicated by the extensive network of αsyn positive fibers at the striatal level (**Figure 1B**). Hαsyn is observed along the axons from neurons of the SN, terminating in the STR. Expression of hαsyn is detected from the most anterior to the most posterior extent of the STR. This event seems to happen rapidly after the viral injection since expression of the transgenes in the terminals is observed after 1 week. (**Figure 1B**, injected side). Moreover, these data demonstrate that the luciferase reporter does not affect the ability of hαsyn protein to be transported along the nigrostriatal pathway.

Because the substrate for hGluc, coelenterazine, does not permeate the blood brain barrier, in vivo luciferase signal was measured in homogenates ex vivo from the SN and STR overexpressing tagged hαsyn at 1 week (n = 7) and at 4 weeks (n = 8) post injection. In each group, all brains were processed at the same time and under the same conditions. During homogenization no detergent, no intense shaking, and no sonication are used. The tissue is briefly homogenized in PBS with a tissue grinder pestle and vortexed for 1 min. The contralateral non-injected sides (STR and SN) were processed in the same manner and served as internal controls for nonspecific background signal. Interestingly, luciferase activity could be detected in the injected SN as early as 1 week post injection (**Figure 2A**), increasing dramatically by 13-fold at 4 weeks (**Figure 2A**). By contrast, in the STR, no luciferase activity is detected at 1 week post injection (**Figure 2B**) whereas, abundant luciferase activity is detected at 4 weeks (**Figure 2B**). The luciferase signal in tissue homogenate from injected animals indicates in vivo luciferase complementation, the surrogate marker of αsyn-αsyn interactions. As expected, in animals injected with only one half (AAV-SL2 only), no luciferase activity was detected after 4 weeks (**Supplementary Figures 1A,B**) demonstrating that fragmented luciferase does not give rise to aberrant luminescence signal. A second control incorporated into our experimental paradigm was a group of animals injected with one half of the complementation pair (AAV-SL1) in the left SN

and the other half (AAV-SL2) in the right SN. After 4 weeks, homogenates from left and right SN or STR were mixed together and assayed for luciferase activity. As expected, no luciferase activity was detected (**Supplementary Figure 1C**) demonstrating that αsyn-αsyn interactions occur in vivo and not ex-vivo during tissue processing.

Reconstitution of luciferase activity in vivo demonstrates the successful complementation of luciferase halves via the formation of αsyn oligomers. Importantly, our data show the presence of αsyn oligomers in the SN as early as 1 week post-delivery of AAV-SL1 and AAV-SL2 (**Figure 2A**). The total level of hαsyn in the SN of the animals was quantified by ELISA at 1 and 4 weeks (**Figure 2C**). No significant difference was detected in hαsyn levels 1 and 4 weeks (**Figure 2C**, 1 week = 0.90 ± 0.05 4 weeks = 0.88 ± 0.22, p > 0.05). Interestingly, although equivalent levels of hαsyn was detected in SN at 1 and 4 weeks, the oligomeric profile was very different (**Figure 2A**). No hαsyn oligomers were detected in the STR at 1 week (**Figure 2B**) despite abundant expression of αsyn by ELISA quantification (**Figure 2D**). There are two ways these data can be interpreted; αsyn oligomers form in the SN and are then transported to the striatal terminals which takes longer than a week or that oligomers form at the terminals over a longer period of time (**Figure 2B**). Regardless, the data support the fact that subcellular location affects αsyn oligomerization kinetics.

### SL1 and SL2 Overexpression Induce Progressive Nigral and Striatal Dopaminergic Neurodegeneration

Previous viral vector rodent models of αsyn overexpression have described a progressive nigral cell loss and striatal terminal neurodegeneration (Kirik et al., 2002; Klein et al., 2002). To confirm that our in vivo αsyn-αsyn bioluminescent complementation model exhibits similar characteristics, we immunostained coronal sections at the level of the SN and STR with an antibody against tyrosine hydroxylase (TH), and performed unbiased stereology and densitometry analysis. At 1 week, TH immunostaining and stereology of the SN indicated no loss of TH+ neurons in the injected side overexpressing SL1 and SL2 when compared to the noninjected side (**Figure 3A**). However, by 4 weeks a general reduction in the dendritic arborization in the SN of the animals was observed (**Figure 3A**) with stereological quantification confirming a 38.5 ± 7.5% dopaminergic cell loss in animals expressing SL1 and SL2 (**Figure 3B**). Also, comparison of nigral

neurons at 1 and 4 weeks revealed a significant difference in the extent of cell loss between the groups. Dopaminergic cell death was further corroborated by the loss of fibers in the STR (**Figures 3C,D**). Densitometric analysis of TH stained sections showed a dense innervation throughout the STR of the non-injected side at all-time points. This was also true of the injected side at 1 week (**Figures 3C,D** OD black bar = 96.9 ± 2.2%). By contrast, at 4 weeks the injected side had a significant reduction of striatal TH+ fibers of 27.7 ± 7.4% relative to the contralateral side (**Figure 3D**). Thus, these data demonstrate a progressive cell and terminal loss through 4 weeks.

### Modulation of αSyn Oligomerization in the Rodent Bioluminescent PCA Model

The fact that αsyn oligomers can be detected in the SN as soon as 1 week post virus delivery make this a very attractive model for rapid screening of modulators of αsyn oligomers. Therefore, we performed a proof-of-concept experiment whereby rats were treated with a previously described heat shock protein inhibitor, SNX-9114 (McFarland et al., 2014). In vitro αsyn-induced aggregation and toxicity is prevented by inhibition of Hsp90 (Klucken et al., 2004; McLean et al., 2004; Danzer et al., 2011; Jones et al., 2014). Hsp90 inhibition results in activation of heat shock factor 1 and by consequence an up-regulation of protective stress-induced proteins such as Hsp70. We recently showed a novel class of Hsp90 inhibitors that could significantly reduce αsyn oligomerization in a cellular model of αsyn aggregation (Putcha et al., 2010; McFarland et al., 2014).

Here we chose to use the Hsp90 inhibitor to validate our 1 week oligomers model. Because SNX-9114 has not been previously validated in vitro, we first evaluated its ability to reduce αsyn oligomer formation in our in vitro complementation assay. Stable cells expressing SL1 and SL2 were plated in a 96 well plate format and treated with 0.01% DMSO or increasing concentrations of the SNX-9114 (25, 50, 100, and 200 nM). Twenty-four hours later, cells were assayed for luciferase activity to determine the effect of the inhibitor on αsyn oligomerization as measured by reconstituted luciferase activity. A decrease in luciferase activity was first detected at 25 nM and 50 nM (**Figure 4A**, p > 0.05), however, at higher doses (100 and 200 nM) SNX-9114 significantly reduced αsyn oligomerization by 32.48 ± 2 and 35.1 ± 4.2% respectively compared to DMSO (**Figure 4A**, p < 0.05). Biochemical analysis of cells expressing SL1SL2 and treated with SNX-9114 confirmed the up-regulation of Hsp70 in a dose-dependent manner (**Figure 4B**) with little to no effect on steady state levels of αsyn except at the highest doses (**Figure 4B**).

To determine if Hsp90 inhibition could reduce αsyn oligomer formation in vivo after 1 week, we stereotactically injected AAV-SL1 and AAV-SL2 into rat SN as described previously. Fortyeight hours after surgery we orally gavaged the rats with 2 mg/kg SNX-9114 or vehicle control. A second gavage of 2 mg/kg SNX-9114 or vehicle was performed 2 days later, and the rats were sacrificed at 7 days post-surgery (**Figure 5A**) A control group of rats transduced with AAV-hGluc were included to control for drug effects on the luciferase enzymatic reaction.

Western blot of homogenates prepared from the SN of injected rats confirmed the induction of Hsp70 in animals treated with SNX-9114 compared to animals treated with vehicle (**Figure 5B**). Furthermore, a luciferase activity assay on the same homogenates demonstrated a significant decrease in αsyn oligomers in the group of animals receiving the Hsp90 inhibitor (**Figure 5C**). No significant difference in hαsyn protein levels was detected between the drug and vehicle control groups when measured by ELISA (**Figure 5D**, p > 0.05). Of note, consistent with our in vitro experiments (**Figure 4B**), SNX-9114 did not directly affect the enzymatic luciferase signal in vivo (**Figure 5E**). These findings suggest that the decreased luciferase signal in the Hsp90 inhibitor treated group is indicative of less αsyn oligomer formation (**Figure 5C**).

#### DISCUSSION

Neuropathologic and genetic studies have provided strong evidence for the involvement of aggregated αsyn in the pathogenesis of synucleopathies (Tu et al., 1998; Dickson et al., 1999; Narhi et al., 1999; Masliah et al., 2000). Our present study proposes a rapid model for tracking αsyn oligomerization in selected brain regions of the rat brain using an ex vivo bioluminescent PCA.

Herein we demonstrate that, our split hGluc AAV-SL1 and SL2 vectors can be utilized to track αsyn oligomerization in vivo. As early as 1 week, we observe the expression of the transgenes along the nigrostriatal pathway and αsyn oligomers in the SN (**Figures 1**, **2**). At this time point no toxicity is detected in the SN and no loss of dopaminergic fibers at the striatal level (**Figure 3**). The expression of hαsyn and presence of oligomers in SN but no cell loss as early as 1 week post-transduction make this an attractive model with which to screen for novel oligomerization modulators. Because no significant difference was detected in total αsyn levels between SN and STR at 1 week, the absence of αsyn oligomers in STR suggests distinct dynamics of oligomer formation in striatum compared to SN or alludes to oligomer formation occurring in the soma and subsequent trafficking of oligomers to the terminals where additional oligomer formation is seeded. Differentiating between these two mechanisms will require further investigation.

By 4 weeks, abundant αsyn oligomers are detected in both SN and STR and cell loss is detected in the SN, with terminal loss detected in the STR. αSyn expression in the SN and STR did not increase even though there was a 13-fold increase in luciferase activity. These data could support a hypothesis whereby increased asyn oligomers correlate to neurotoxicity, however further studies will be required before this can be confirmed.

Given the essential role of αsyn oligomers in the pathogenesis of synucleopathies it is essential that we develop reliable methods for their detection.  They also hold potential as therapies and could be a relevant disease biomarkers. However, such antibodies cannot monitor the oligomeric state of αsyn in a live cell environment. PCA strategies using fluorescence or bioluminescence reporters have been developed and have proven to be a powerful approach to study protein aggregation. Recently, we generated a rodent model using YFP venus PCA to monitor αsyn oligomerization in live animals (Dimant et al., 2014). Using fluorescence as readout, we directly detected αsyn oligomers and monitored αsyn aggregation in cortical neurons of living mice using two photon microscopy. Recently, a mouse model expressing firefly luciferase

tagged-αsyn in vivo was generated and bioluminescent imaging (IVIS) was used to non-invasively capture αsyn oligomerization in the mouse brain (Aelvoet et al., 2014). Both of these in vivo models have distinct advantages such as the ability to follow the response in the same animals over time, providing a kinetic readout of the oligomerization process. In our model we measure the oligomerization ex-vivo. Although, all brains were processed at the same time and under the same conditions, we cannot exclude the possibility that the extraction procedure could perturb αsyn aggregation state. On the other hand, even though in vivo imaging represents a good tool to monitor αsyn, these methods require expensive instrumentation, are time-consuming, and involve anesthesia and systemic substrate injections in the case of the firefly luciferase PCA. Lastly, the two photon microscopy is limited by photo absorption of the tissue and does not allow deep brain structure imaging.

The model presented herein has the advantage of having a rapid readout without expensive equipment. HGluc may also provide increased sensitivity as it is over 100-fold more sensitive than the commonly used luciferases, Firefly, and Renilla reniformis (Tannous et al., 2005; Remy and Michnick, 2006). Also Gluc is a smaller reporter (185 amino acid; Tannous et al., 2005) compared to the other forms of luciferase or the fluorescence reporter. Finally, hGluc PCA is a fully reversible interaction unlike the assay based on YFP venus fluorophore, thus facilitating the detection of the kinetics of protein complex assembly/disassembly in vivo. Better spatial and temporal resolution can be achieved compared with in vivo imaging and we believe that hGluc PCA may be more efficient and effective for some approaches. In our current experiment, oligomers were quantified ex vivo for several reasons. Brain lysate analysis provides a more accurate semi quantitative measure of aggregates and most importantly, allows a combination of different analyses on the same sample (e.g., ELISA, WB) for each brain. Because the PCA assay is effective in tissue directly placed in a micro titer plate, its utility to measure oligomers in in vivo brain slice preparations is also a possibility. Also, it is well established that αsyn oligomers can be transmitted from neuron(s) to neuron(s), thus inducing pathology along neuroanatomical pathways in the brain (El-Agnaf et al., 2006; Emmanouilidou et al., 2010). It remains to be determined if αsyn induces neurodegeneration via intracellular- or extracellular-associated mechanisms. Our AAVmodel, coupled with in vivo microdialysis, could be a useful tool for the detection and analysis of αsyn oligomers in the interstitial fluid (ISF) of living animals, allowing a real time monitoring of in vivo processes and opening possibilities for αsyn transmission studies. One disadvantage is that coelenterazine, the substrate of hGluc, does not cross the blood brain barrier and therefore excludes the use of our model for imaging in live animals. Despite this limitation, the use of hGluc PCA in vivo provides a rapid model to track αsyn oligomers depending on the desired readout.

To investigate the potential of our in vivo bioluminescent PCA to identify compounds that interfere with αsyn-αsyn interaction or modulate oligomerization, in a little as 1 week, we administered a cohort of AAV SL1 and SL2 transduced rats with 2 mg/kg SNX-9114, for one week. Hsp90 inhibition has been shown to modulate αsyn aggregation and toxicity (Auluck et al., 2002; Kalia et al., 2010; Luo et al., 2010; Putcha et al., 2010) and we have previously established that pharmacological pretreatment with geldanamycin, a naturally occurring Hsp90 inhibitor, protects against αsyn-induced toxicity and leads to degradation of high molecular-weight species (McLean et al., 2004) More recently we described the efficacy of novel brain permeable Hsp90 inhibitors to reduce the formation of αsyn oligomeric species in a dose dependent manner using our bioluminescent PCA system in H4 neuroglioma cells (Putcha et al., 2010). In line with these previous data, we show here that an Hsp90 inhibition can inhibit αsyn oligomerization in our AAV-SL1 and AAV-SL2 rodent model, and most importantly, we demonstrated that our luciferase assay is sensitive enough to detect these changes. In addition it is noteworthy to mention that the level of oligomers was assessed as early as 1 week after the AAV-injection, at a time when we do not detect cell death (**Figure 3B**) but we have an abundance of oligomers in the SN (**Figure 2A**). Therefore, the detected decrease in luciferase activity at 1 week in SN in animals receiving Hsp90 inhibitor, can be directly attributed to a decrease of αsyn oligomers.

While several strategies have been developed to monitor the modulation of αsyn aggregation in vivo and in vitro, the major advantage of the present approach over other techniques is the ability to quantify a change in αsyn oligomerization in a short period of time (1 week) with a fast, simple, and sensitive assay. In conclusion, our bioluminescent in vivo model represents a

#### powerful new tool to study spatial and temporal changes in αsyn aggregation in response to new therapeutic agents that modulate αsyn oligomerization.

### AUTHOR CONTRIBUTIONS

MD carried out the study design, animal procedures, histology, analysis, and drafting of the manuscript. TT performed immunohistochemistry, stereology analysis, densitometry analysis, and participated in all animal procedures. DJ, participated in the study design, animal procedures, and editing of the manuscript. SM and AB performed the in vitro assay. MY, assisted with animal procedures. WH provided the compound SNX-9114. PM participated in the experimental design, coordination, interpretation, drafting, and editing of the manuscript. All authors read and approved the final manuscript.

#### ACKNOWLEDGMENTS

We thank the Mayo Clinic Viral Vector core, Ms. Karen Jansen-West, Ms. Ena Whitelaw, and Ms Emilie Perkerson for high titer AAV production. This work was funded by NIH NS073740 to PM.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fnins. 2015.00511

Supplementary Figure 1 | Luciferase activity in control rats. (A) Animals were stereotactically injected with one viral vector (AAV8-SL2, *n* = 2). After 4 weeks, the luciferase activity was measured. No specific signal was detected in the injected side since the luciferase activity was similar to the non-injected side. In both regions, SN (A) and STR (B,C) To verify that the complementation occurs *in vivo* and not during homogenization process, animals were injected with AAV8-SL1 In the left SN and with AAV-SL2 in the right SN (*n* = 2). At 4 weeks homogenate of SN left and right or STR left and right, were mixed together respectively and luciferase activity was measured. No activity was detected in the mixture of SN or STR.

### REFERENCES

familial Parkinson's disease. Lancet 364, 1167–1169. doi: 10.1016/S0140- 6736(04)17103-1

overexpression of alpha-synuclein in the nigrostriatal system. J. Neurosci. 22, 2780–2791.

system atrophy brains contain insoluble alpha-synuclein. Ann. Neurol. 44, 415–422. doi: 10.1002/ana.410440324

Winner, B., Jappelli, R., Maji, S. K., Desplats, P. A., Boyer, L., Aigner, S., et al. (2011). In vivo demonstration that alpha-synuclein oligomers are toxic. Proc. Natl. Acad. Sci. U.S.A. 108, 4194–4199. doi: 10.1073/pnas.1100976108

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Delenclos, Trendafilova, Jones, Moussaud, Baine, Yue, Hirst and McLean. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Is Alpha-Synuclein Loss-of-Function a Contributor to Parkinsonian Pathology? Evidence from Non-human Primates

Timothy J. Collier 1, 2, D. Eugene Redmond Jr. 3, 4 , Kathy Steece-Collier 1, 2, Jack W. Lipton1, 2 and Fredric P. Manfredsson1, 2 \*

*1 Department of Translational Science & Molecular Medicine, Michigan State University, Grand Rapids, MI, USA, 2 Hauenstein Neuroscience Center, Mercy Health Saint Mary's, Grand Rapids, MI, USA, 3 Departments of Psychiatry & Neurosurgery, Yale University School of Medicine, New Haven, CT, USA, 4 Axion Research Foundation, Hamden, CT, USA*

#### Edited by:

*Mark R. Cookson, National Institutes of Health, USA*

#### Reviewed by:

*Luke Esposito, ProtaMed, Inc., USA Claudia Manzoni, University of Reading, UK*

#### \*Correspondence:

*Fredric P. Manfredsson<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Neurodegeneration, a section of the journal Frontiers in Neuroscience*

Received: *02 December 2015* Accepted: *11 January 2016* Published: *29 January 2016*

#### Citation:

*Collier TJ, Redmond DE Jr., Steece-Collier K, Lipton JW and Manfredsson FP (2016) Is Alpha-Synuclein Loss-of-Function a Contributor to Parkinsonian Pathology? Evidence from Non-human Primates. Front. Neurosci. 10:12. doi: 10.3389/fnins.2016.00012*

Accumulation of alpha-synuclein (α-syn) in Lewy bodies and neurites of midbrain dopamine neurons is diagnostic for Parkinson's disease (PD), leading to the proposal that PD is a toxic gain-of-function synucleinopathy. Here we discuss the alternative viewpoint that α-syn displacement from synapses by misfolding and aggregation results in a toxic loss-of-function. In support of this hypothesis we provide evidence from our pilot study demonstrating that knockdown of endogenous α-syn in dopamine neurons of non-human primates reproduces the pattern of nigrostriatal degeneration characteristic of PD.

Keywords: alpha-synuclein, Parkinson's disease, non-human primates, substantia nigra, dopamine

## ALPHA-SYNUCLEIN IN PARKINSON'S DISEASE

The best-validated participant in the molecular pathology of Parkinson's disease (PD) is alphasynuclein (α-syn). Mutations and multiplications of the α-syn SNCA gene locus both produce inherited forms of PD (Polymeropoulos et al., 1997; Singleton et al., 2003; Ibanez et al., 2004). In addition, the presence of α-syn in Lewy bodies and neurites of midbrain dopamine (DA) neurons, the histological hallmark of PD, provides evidence for its association with idiopathic PD (Spillantini et al., 1997). While a pervasive viewpoint postulates that excess α-syn and consequent aggregation directly fuels neurotoxicity in a toxic gain-of-function event, it also is acknowledged that α-syn aggregation may endanger neurons by removing the protein from its normal cellular location and diminishing its function in a toxic loss-of-function event (Perez and Hastings, 2004; Cookson, 2006; Kanaan and Manfredsson, 2012). This issue remains a topic of debate.

Despite this ambiguity, approaches that may indiscriminately reduce α-syn in the central nervous system represent an active area of research as an approach for treating PD (e.g., AFFiRiS PD01, Prothena/Roche PRX002). In this Hypothesis and Theory article, we discuss evidence from rodent and non-human primate experiments suggesting that Parkinson's-like degeneration of the nigrostriatal DA system can be reproduced by eliminating endogenous α-syn from DA neurons vulnerable to degeneration in PD. These studies support the tentative conclusion that α-syn elimination therapies that do not distinguish between native and abnormal forms may compromise the viability of DA neurons and should proceed with caution.

#### Collier et al. Alpha-Synuclein Loss-of-Function

### KNOCKDOWN OF α-SYN IN THE ADULT RAT RESULTS IN DOSE-DEPENDENT NIGROSTRIATAL DEGENERATION

The lack of overt pathology in α-syn germ-line knockout animals (Abeliovich et al., 2000) and the commonly proposed toxic gain-of-function of the α-syn mutant versions (A53T, A30P, E46K; Kruger et al., 1998; Giasson et al., 2002; Klein et al., 2002; Zarranz et al., 2004) led many investigators, us among them, to pursue targeted knockdown of α-syn expression as a potential therapeutic avenue for PD.  In our rodent studies, neurodegeneration could be rescued by coexpression of rat α-syn (rendered insensitive to the shRNA) demonstrating that neuronal loss was explicitly due to a toxic loss-of function of α-syn and not due to non-specific shRNAmediated toxicity. Moreover, by utilizing several distinct shRNA sequences displaying varying efficiencies of endogenous rat α-syn mRNA knockdown we showed that the extent of neuronal loss was dependent on the level of remaining α-syn. The ability to rescue neurons by co-expression of rat α-syn and the fact that toxicity was proportional to the efficiencies of shRNAs demonstrate that neuronal loss was not due to "off-targeting" of other endogenous mRNAs or due to non-specific shRNA toxicity.

## KNOCKDOWN OF α-SYN IN THE NON-HUMAN PRIMATE RESULTS IN DOSE-DEPENDENT NIGROSTRIATAL DEGENERATION: A CASE STUDY

To begin to further characterize the consequences of knockdown of endogenous α-syn and determine whether effects seen in rats were reproducible in a species more closely related to humans, we generated an α-syn shRNA specific for St.Kitts green monkeys (Chlorocebus sabaeus). We injected α-syn shRNA or scrambled shRNA, of two different titers, into the SN of individual monkeys (N = 4 total), waited 3 months, and examined DA neuron numbers, morphology, striatal innervation, and striatal DA content. α-Syn shRNA produced region-specific, titer-related, degeneration of SN tyrosine hydroxylase-positive (TH+) neurons and innervation of the striatum, reproducing the pattern of nigrostriatal degeneration observed in PD: SN degeneration was exaggerated in ventral tier neurons (vtSN; Gibb and Lees, 1991) with relative sparing of adjacent ventral tegmental area (VTA) DA neurons, and loss of TH+ fibers in the putamen (Pt) exceeded denervation of the caudate nucleus (Cd) (Kish et al., 1988; **Figure 1**). Stereologic quantification of TH+ neurons confirmed the qualitative observations: loss of TH+ neurons was greatest in vtSN with the VTA exhibiting relatively little neuron loss. However, the general pattern of TH+ neuron loss was not different in the High and Low titer α-syn shRNA conditions. These neurons also contain neuromelanin, and when counts of neuromelanin-only positive cells were added to the analysis, titer-related differences emerged. The Low titer shRNA condition was associated with the preservation of significantly more neuromelanin-only neurons, suggesting that while loss of TH+ phenotype was equivalent across titers, overt loss of neurons was greater in the High titer condition. The presence of TH-negative, neuromelanin-positive neurons is suggestive of ongoing pathology, an observation that also is seen in early PD. Importantly, co-localization of green fluorescent protein (GFP) as a marker of viral transduction within surviving midbrain DA neurons confirmed that dorsal SN and VTA neurons were transduced, but showed less degeneration (**Figure 1**). Titerrelated differences in striatal DA depletion also were detected. Deficits in DA in the caudate nucleus and putamen were exaggerated in the α-syn shRNA High titer condition and a significant increase in the homovanillic acid (HVA)/DA ratio was only detected in the High titer subject. Increase in the HVA/DA ratio is known to be associated with a compensatory response to ongoing significant degeneration of DA neurons (Zigmond et al., 1990).

## HOW DOES LOSS OF PRESYNAPTIC α-SYN CONTRIBUTE TO NEURODEGENERATION?

The scope of our one subject per treatment condition pilot study in non-human primates has clear limitations. Nevertheless, the results confirm our previous findings in the rat (Gorbatyuk et al., 2010; Kanaan and Manfredsson, 2012). At this stage our findings, while internally consistent among the treatment conditions, must be considered as generating a hypothesis, rather than being conclusive. While the idea that maintaining α-syn expression is critical for survival of populations of adult neurons has received relatively little investigation, the high prevalence and widespread distribution of α-syn in brain inarguably reflects an important function in synapses. While direct evidence for the loss-offunction hypothesis is in its early stages, many studies provide indirect support for the concept.

Alpha-synuclein exhibits a dramatic redistribution within the neuronal compartment as a function of aging: the best-validated primary risk factor for PD. A quantitative morphological postmortem analysis of α-syn immunoreactivity within SN DA neurons of humans reveals a distinct pattern of increased staining within neuronal soma with advancing chronological age (Chu and Kordower, 2007). Samples were from individuals with a mean age of 31 years (young group, N = 6), 55 years (middleaged, N = 4) and 84 years (aged, N = 8). Alpha-synuclein immunoreactivity was rarely detected in the soma of SN neurons in young individuals and was restricted to its normal location in the neuropil. With advancing age, a progressive increase in somatic immunoreactivity was observed, culminating in a 639% increase in aged subjects. Quantitation of the intensity of somatic staining on a per neuron basis confirmed this pattern, illustrating a mean 57% increase in aged subjects as compared to young. It is noteworthy that none of these samples exhibited α-syn aggregates, suggesting that this intracellular redistribution of α-syn may be an ongoing aging-related event that precedes

processes of aggregation. The same pattern was observed in aging monkeys. Of importance to the loss-of-function hypothesis, no comparison to α-syn distribution/levels in striatum was performed, providing no opportunity to assess whether increased somatic levels are associated with decreased synaptic levels.

Oxidative stress is an established threat to DA neuron viability and can be significantly exacerbated by DA mishandling (i.e., increased cytosolic DA). Indeed, multiple lines of evidence suggest a regulatory role for synuclein in dopamine neurotransmission (Perez et al., 2002; Maries et al., 2003; Wersinger et al., 2003), consistent with the view that loss of presynaptic α-syn may contribute to increased oxidative stress via dysregulation of DA biosynthesis, release and metabolism. Studies in primary cultures of DA neurons demonstrate that in conditions of elevated oxidative stress, endogenous levels of α-syn increase and provide neuroprotection (Quilty et al., 2006; Musgrove et al., 2011). Similarly, transgenic mice over-expressing α-syn are resistant to paraquat-induced toxicity for DA neurons known to be associated with oxidative stress (Manning-Bog et al., 2003). Thus, endogenous α-syn likely provides an important DA neuron defense against oxidative damage.

Mishandling of synaptic DA and the toxic byproducts of its metabolism in the cytosol have a long history of association with the vulnerability of these neurons (e.g., Perez and Hastings, 2004; Segura-Aguilar et al., 2014). Studies of transgenic mice deficient in α-syn, while free of overt DA neuron pathology in adulthood, exhibit changes in DA neurotransmission, with some of these becoming exaggerated in aged animals. Adult, triple knock-out mice for all synuclein family proteins exhibit elevated evoked release of DA in striatum, enhanced turnover and reduced presynaptic DA stores (Anwar et al., 2011). Study of two α-syn deficient mouse lines found evidence for increased stimulated DA overflow in striatum, higher basal extracellular DA levels, decreased expression of the dopamine transporter (DAT) and reduced DA reuptake (Chadchankar et al., 2011). Aged (24– 26 months old) α-syn null mice exhibit significant reduction of striatal DA, a decrease in TH+ fibers and decreased striatal levels of TH and DAT (Al-Wandi et al., 2010). These data support the view that α-syn is a pivotal presynaptic regulator of DA neurotransmission and that disruption of this process may result in chronic accumulation of DA in the cytosol.

Perhaps the most direct evidence for an α-syn loss-of-function process related to aggregation comes from the mouse pre-formed fibril (PFF) model of synucleinopathy (Luk et al., 2012). In this model, intracerebral injection of pre-formed fibrils of α-syn seeds misfolding of endogenous α-syn, resulting in progressive accumulation of cytoplasmic aggregates of the protein. In a recent report (Osterberg et al., 2015) the formation of aggregates was tracked over time. The findings indicate that as inclusions mature into a compact form, detection of soluble α-syn shows a parallel decline.

### IS TOXICITY DUE TO LOSS OF α-SYN RELEVANT TO PARKINSON'S DISEASE ETIOLOGY?

Direct assessment of the relevance of the α-syn loss-of-function hypothesis to PD is a challenge. At present, there is no imaging protocol specific for α-syn, although this is an active area of research. To the best of our knowledge, there has been no study of striatal synaptosome preparations comparing PD to controls and there may be significant technical limitations to this approach with postmortem tissue. In addition, since numerous neuronal circuits project to the striatum, it may be difficult to discern what effect is directly attributable to nigrostriatal projections. Studies of α-syn in body fluids as a biomarker for PD may hold clues, but often are difficult to interpret in the context of entry of α-syn into these compartments and the potential presence or impairment of mechanisms operating to maintain homeostasis. However, α-syn is enriched in brain and blood, and from the context of PD as a "whole body" syndrome it can be argued that a systemic change in α-syn that characterizes PD may be detectable in blood as a reflection of a general process that affects brain. Taken together, biomarker studies targeting α-syn present a set of mixed results (Malek et al., 2014 for review), documenting increases, decreases and no differences in PD subjects as compared to controls. Many of these conclusions likely are linked to differences in sample collection, preparation and the assay employed. One recent report takes a different approach to circumvent the complexities of accurately measuring α-syn protein in blood and measured αsyn transcripts. They analyzed blood samples from three large cohorts of PD patients and controls from prior and ongoing clinical trials individually. All three cohorts supported the finding of an approximate 20% reduction in α-syn transcripts even in newly diagnosed PD patients (Locascio et al., 2015). However, this finding could be interpreted as support of loss-of-function, gain-of-function, or be unrelated to processes operating in brain.

Abundant information from human genetic studies document a clear association between increases in α-syn gene-dosing, or mutations, that exacerbate aggregation and lead to PD. In line with our hypothesis, these all are processes that can directly increase the sequestration of α-syn away from the synapse via increased aggregation kinetics (e.g., enhanced molecular crowding). Nevertheless, one would expect that if α-syn is crucial for maintaining the viability of DA neurons, human genetic data on risk of PD also would implicate mutations or polymorphisms that result in decreased α-syn. Indeed, a recent report has described exactly that (Markopoulou et al., 2014). The dinucleotide repeat REP1 is positioned upstream of the SNCA transcription start site. Polymorphisms at this site are associated with either increased or decreased SNCA expression. Contrary to the investigator's hypothesis that polymorphisms associated with decreased expression of α-syn would be associated with decreased risk of PD, these low α-syn individuals exhibited increased risk of developing PD as well as worse motor and cognitive outcomes.

### LOSS-OF-FUNCTION, GAIN-OF-FUNCTION OR A BIT OF BOTH?

Loss-of-function and gain-of-function hypotheses are not irreconcilable. The absence of an overt parkinsonian phenotype and DA neuron degeneration in germline α-syn knockout mice does not directly address the loss-of-function hypothesis. Whole genome expression analysis of SNCA(−/−) mice identified differential expression of 369 transcripts as compared to wildtype animals (Kuhn et al., 2007). This includes increased expression of transcripts of other synuclein family members, 14-3-3 proteins, TH and neurotrophic factors, decreased expression of pro-apoptotic transcripts, and changes in genes directly involved in vesicle function and neurotransmission. Thus, the lack of a severe phenotype in these animals is likely the result of complex germline compensation for α-syn function. While heritable forms of PD associated with duplication and triplication of the SNCA gene arguably support the toxic gainof-function hypothesis, germline α-syn overexpression mice do not directly support this contention. The available mouse models exhibit little or no degeneration of SN DA neurons, although similar to α-syn knockout mice, show prominent changes in striatal DA neurotransmission (e.g., Chesselet and Richter, 2011; Crabtree and Zhang, 2012 for reviews). Nevertheless, if the α-syn transgene is induced in the mature animal, nigral degeneration does occur (Lin et al., 2012). Thus, it is plausible that germline compensation, similar to that seen in the knockout, occurs in overexpression transgenic models as well.

While germline transgenic models do little to inform the toxic gain-of-function/loss-of-function debate it is inarguable that too much α-syn and mutant α-syn can produce direct neurotoxicity. In vitro studies demonstrate that specific conformations of αsyn exert cytotoxic effects by permeabilizing vesicles, altering calcium flux, impairing mitochondrial function, and inducing apoptosis (Volles et al., 2001; Pieri et al., 2012; Luth et al., 2014; Pacheco et al., 2015). In addition, viral vector-mediated overexpression of α-syn in DA neurons uniformly produces significant, albeit variable in magnitude, neurodegeneration (e.g., Van der Perren et al., 2014 for review). However, in this case it is unclear what precipitates neurodegeneration. Nonetheless, in line with our hypothesis, and with data gained from the PFF model (i.e., Osterberg et al., 2015) it is plausible that loss of

soluble endogenous α-syn to aggregation is one contributor to neurotoxicity. Thus, experimental support for both viewpoints suggests a "Goldilocks" biology commonly exhibited by many molecules: too much is bad, too little is bad, enough is just right (Kanaan and Manfredsson, 2012).

## CONCLUSIONS

In this Hypothesis and Theory article we address the alphasynuclein gain-of-function vs. loss-of-function debate as it applies to DA neuron pathology in PD. Our experimental findings in rats and non-human primates, combined with support for the concept from other reports in the literature, leads to the proposal that loss-of-function plays a significant role in this pathology. On its surface, the α-syn loss-of-function hypothesis seems counterintuitive since α-syn accumulation as aggregates, in the form of Lewy bodies, is a key neuropathologic feature of PD. However, while experimental observations largely have been interpreted as suggestive of a cytotoxic role for α-syn, it does not preclude the alternate interpretation that α-syn aggregation reflects processes that prevent α-syn from performing its normal biological functions. Our recent findings in non-human primates support the contention that α-syn loss-of-function in midbrain DA neurons accurately reproduces the pattern of nigrostriatal degeneration observed in PD, and that α-syn aggregation, acting as a "sink," is one possible driver of this process rather than a direct toxic culprit in cell loss (**Figure 2**). The findings suggest reconsideration of the relationship of α-syn to PD pathogenesis and caution in the implementation of α-syn clearance therapeutic strategies that do not distinguish between natural forms and pathological forms.

## AUTHOR CONTRIBUTIONS

TC conception, design, acquisition analysis, data interpretation, drafting and revision, DR design, acquisition analysis, revision, KC acquisition analysis, revision, JL acquisition analysis, data interpretation, revision, FM conception, design, acquisition analysis, data interpretation, drafting and revision.

## ACKNOWLEDGMENTS

The authors are grateful for the significant technical contributions of Allyson Cole-Strauss, Brian Daley, Nathan Kuhn, Nathan Marckini, Zachary Mattingly, Ernell Nisbett, Xavier G. Morton, and Ryan M. Malpass.

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some clinical and basic implications. Trends Neurosci. 13, 290–296. doi: 10.1016/0166-2236(90)90112-N

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Collier, Redmond, Steece-Collier, Lipton and Manfredsson. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Altered Expression Patterns of Inflammation-Associated and Trophic Molecules in Substantia Nigra and Striatum Brain Samples from Parkinson's Disease, Incidental Lewy Body Disease and Normal Control Cases

Douglas G. Walker 1, 2 \*, Lih-Fen Lue1, 2, Geidy Serrano1 , Charles H. Adler 3 , John N. Caviness 3 , Lucia I. Sue1 and Thomas G. Beach1

#### Edited by:

*Timothy J. Collier, Michigan State University, USA*

#### Reviewed by:

*Steve M. Gentleman, Imperial College London, UK Varun Kesherwani, University of Nebraska Medical Center, USA*

> \*Correspondence: *Douglas G. Walker<EMAIL_ADDRESS>
> > Specialty section:

*This article was submitted to Neurodegeneration, a section of the journal Frontiers in Neuroscience*

Received: *02 October 2015* Accepted: *21 December 2015* Published: *14 January 2016*

#### Citation:

*Walker DG, Lue L-F, Serrano G, Adler CH, Caviness JN, Sue LI and Beach TG (2016) Altered Expression Patterns of Inflammation-Associated and Trophic Molecules in Substantia Nigra and Striatum Brain Samples from Parkinson's Disease, Incidental Lewy Body Disease and Normal Control Cases. Front. Neurosci. 9:507. doi: 10.3389/fnins.2015.00507* *1 Banner Sun Health Research Institute, Sun City, AZ, USA, 2 Neurodegenerative Disease Research Center, Biodesign Institute, Arizona State University, Tempe, AZ, USA, 3 Neurology, Mayo Clinic College of Medicine, Scottsdale, AZ, USA*

Evidence of inflammation has been consistently associated with pathology in Parkinson's disease (PD)-affected brains, and has been suggested as a causative factor. Dopaminergic neurons in the substantia nigra (SN) pars compacta, whose loss results in the clinical symptoms associated with PD, are particularly susceptible to inflammatory damage and oxidative stress. Inflammation in the striatum, where SN dopaminergic neurons project, is also a feature of PD brains. It is not known whether inflammatory changes occur first in striatum or SN. Many animal models of PD have implicated certain inflammatory molecules with dopaminergic cell neuronal loss; however, there have been few studies to validate these findings by measuring the levels of these and other inflammatory factors in human PD brain samples. This study also included samples from incidental Lewy body disease (ILBD) cases, since ILBD is considered a non-symptomatic precursor to PD, with subjects having significant loss of tyrosine hydroxylase-producing neurons. We hypothesized that there may be a progressive change in key inflammatory factors in ILBD samples intermediate between neurologically normal and PD. To address this, we used a quantitative antibody-array platform (Raybiotech-Quantibody arrays) to measure the levels of 160 different inflammation-associated cytokines, chemokines, growth factors, and related molecules in extracts of SN and striatum from clinically and neuropathologically characterized PD, ILBD, and normal control cases. Patterns of changes in inflammation and related molecules were distinctly different between SN and striatum. Our results showed significantly different levels of interleukin (IL)-5, IL-15, monokine induced by gamma interferon, and IL-6 soluble receptor in SN between disease groups. A different panel of 13 proteins with significant changes in striatum, with IL-15 as the common feature, was identified.  The results demonstrate the feasibility of profiling inflammatory molecules using diseased human brain samples, and have provided additional targets to validate in relation to PD pathology.

Keywords: inflammation, pathology, cytokines, antibody array, dopaminergic cell loss, Parkinson's disease, microglia, astrocytes

#### INTRODUCTION

Parkinson's disease (PD) is a chronic, progressive neurodegenerative disorder characterized by symptoms of tremor, bradykinesia, ataxia and rigidity, and is the main cause of movement disorders in the elderly. The current estimates of PD are one million cases in the U.S.A. with 60,000 cases being added each year (Parkinson's Disease Foundaton, 2015). This leads to significant morbidity and mortality in affected individuals along with the considerable cost of healthcare for PD patients. Sufferers of PD also have significant risk of progressing to dementia (Aarsland et al., 2005). There have been major strides in understanding the cause(s) of PD, but there is still lack of consensus of the sequence of events that lead to the loss of the dopaminergic neurons of the substantia nigra (SN) pars compacta and results in loss of dopaminergic neurotransmission in the striatum. Much research on disease mechanisms has focused on the properties of the presynaptic vesicle protein α-synuclein whose modifications can lead to the formation of aggregated and neurotoxic species (Vekrellis and Stefanis, 2012; Lawand et al., 2015; Osterberg et al., 2015; Sian-Hulsmann et al., 2015). Aggregated α-synuclein, particularly the phosphorylated form, is the major component of Lewy bodies, the major pathological feature of PD brains (Fujiwara et al., 2002). Most of the identified mutations in α-synuclein appear to increase the risk of PD by increasing its tendency to aggregate (Polymeropoulos, 1997; Baba et al., 1998; Conway et al., 1998; Ancolio et al., 2000; Ostrerova-Golts et al., 2000; Ghosh et al., 2013; Giráldez-Pérez et al., 2014).

Inflammation has also been a well-established feature of PD pathology (McGeer et al., 1988; Boka et al., 1994; Mogi et al., 1994; Hunot et al., 1999; Brochard et al., 2009), and some data have suggested it could be the primary pathological cause for SN neuronal cell death (Tansey et al., 2007, 2008; Vivekanantham et al., 2015). Inflammation in PD brains could be caused by different factors, but a number of studies have shown that aggregated forms of α-synuclein can activate microglia to produce toxic molecules that contribute to dopaminergic cell death (Zhang et al., 2005; Couch et al., 2011; Béraud et al., 2013; Acosta et al., 2015). Alpha-synuclein activation of microglia can be mediated through the Toll-like receptor (TLR)- 2 (Codolo et al., 2013; Kim et al., 2013; Doorn et al., 2014; Daniele et al., 2015), though other microglial receptors have also been implicated, including TLR-4 (Stefanova et al., 2011; Fellner et al., 2013) and purinergic receptor P2X7 (Jiang et al., 2015). A recent immunohistochemical study of SN from control, ILBD and PD cases for the inflammation marker TLR-2 showed increased numbers of TLR-2-positive microglia in ILBD cases compared to PD cases (Doorn et al., 2014). This suggested that some inflammatory changes could be happening at early stages prior to development of symptoms of PD. By contrast, there was progressive increase from control to PD in numbers of CD68-positive amoeboid microglia/macrophages, a marker associated with phagocytosis, which correlated with an increase in α-synuclein deposits (Doorn et al., 2014). Independent of the presence of pathological α-synuclein, the human SN appears to be particularly sensitive to inflammation, possibly due to higher concentrations of iron, and neuromelanin, both of which can contribute to an environment of enhanced oxidative stress (Hirsch, 1993; Béraud et al., 2013; Taylor et al., 2013; Fischer and Maier, 2015). Purified neuromelanin can directly activate microglia to a proinflammatory state (Wilms et al., 2003; Zhang et al., 2013).

Animal models for PD have provided the most convincing evidence for how inflammation could be directly linked to SN neuronal cell loss. A widely used model involves the administration of the bacterial cell wall extract lipopolysaccharide (LPS) (Couch et al., 2011; Qin et al., 2013; Tanaka et al., 2013; Sharma and Nehru, 2015). Both direct injection of LPS into the SN or by intraperitoneal injection can lead to enhanced inflammation in the brain and selective loss of SN neurons. Administration of LPS results in increased production of free radicals and potentially toxic cytokines, including tumor necrosis factor-α (TNF-α) (McGuire et al., 2001; Pei et al., 2007; Tansey et al., 2007, 2008; Zhao et al., 2007; Gao et al., 2011; Tran et al., 2011; Montgomery and Bowers, 2012; Qin et al., 2013). Another feature of inflammation-induced models of PD is disruption of the blood-brain barrier, which enhances inflammation by permitting influx of components of cell-mediated immunity (Carvey et al., 2005b, 2006; Desai et al., 2007; Monahan et al., 2008).

The spread of abnormal forms of α-synuclein along neuroanatomical pathways is a significant pathological mechanism in humans and animal models of synucleinopathies (Beach et al., 2009; Luk et al., 2012; Masuda-Suzukake et al., 2014; Paumier et al., 2015). In animal models, instrastriatal injection of preformed α-synuclein fibrils resulted in neurodegeneration and inflammation in the SN; the reverse effect occurs with α-synuclein administered into the SN resulting in striatum pathology and inflammation (Koprich et al., 2010; Luk et al., 2012; Osterberg et al., 2015; Paumier et al., 2015). Direct injection of LPS into striatum or SN resulted in degeneration of nigrostriatal pathway neurons and motor impairments, along with microglial activation (Choi et al., 2009; Couch et al., 2011). Co-administration of α-synuclein and LPS can significantly enhance the generation of nigrostriatal pathology (Couch et al., 2011; Gao et al., 2011). As it is still not known if spread of pathology or neuroinflammation occurs from striatum to SN or vice-versa, as part of this study, we sought to compare the changes in inflammation and trophic molecules between SN and striatum with disease progression. Recent mouse models of PD have focused on α-synuclein, either overexpression of normal or mutated α-synuclein under transgene control or overexpression by administration of α-synuclein viral transduction vectors (Watson et al., 2012; Béraud et al., 2013; Gardai et al., 2013; Harms et al., 2013). Animal models of PD can also be developed using the dopaminergic neurotoxin 1-methyl-4 phenyl-1,2,3,6 tetrahydropyridine (MPTP) to lesion the SN and striatum. This model can reproduce many PD features in non-human primates (Ohnuki et al., 2010). This study carried out gene expression profiling of SN and striatum tissue and showed significant downregulation of neuronal and dopaminergic genes in lesioned animals. With respect to inflammation-related genes, only upregulation of glial fibrillary acidic protein (GFAP), interleukin (IL)-11, and chemokines CXCL13 and CXCL4 were detected (Ohnuki et al., 2010).

Although there have been many immunohistological studies that used antibodies to activated microglia to demonstrate inflammation in PD brains, there have been few studies that biochemically measured levels of cytokines or other inflammation-associated molecules in human brain samples (Nagatsu et al., 2000a,b). A recent proteomics analysis comparing SN tissue from PD and control cases identified cytosolic nonspecific dipeptidase 2 to be upregulated in PD tissue, but not inflammatory or growth factor molecules. This may have been due to lack of sensitivity of the methods employed to detect these low-abundance molecules (Licker et al., 2012). Such experiments are technically difficult as only low concentrations of key molecules are present in tissue. Cerebrospinal fluid (CSF), plasma and blood have also been used as surrogates to follow brain changes in PD (Rocha et al., 2014), but these results have generally been inconsistent between studies. Increased levels of IL-2, IL-6, and TNF-α have been detected in CSF samples of PD subjects compared to controls (Mogi et al., 1994, 1996). Screening of sera from control, PD, multiple system atrophy and corticobasal syndrome cases using a similar antibody array as used in this study, showed only platelet-derived growth factor (PDGF)-BB and prolactin having significant disease associated differences (Mahlknecht et al., 2012). Measurement of cytokines in CSF from PD and control cases identified significant changes in the cytokines/growth factors vascular endothelial growth factor (VEGF), placental growth factor (PIGF), soluble VEGF receptor (sVEGFR2), and angiopoietin2 (ANG2), associated with angiogenesis (Janelidze et al., 2015).

To address the problems of sensitivity, studies have used mRNA gene expression profiling of PD SN tissue, or laserdissected SN dopaminergic neurons, to identify disease differences. These techniques have high sensitivity for detecting low-abundance gene expression, but there has been lack of consensus on the PD-associated differentially expressed genes between studies. However, these studies have identified multiple pathways affected in PD, with downregulation of genes associated with synaptic function, cytoskeletal function and neuroprotection and also ubiquitin-proteosome and mitochondrial function genes being features (Grünblatt et al., 2004; Hauser et al., 2005; Mandel et al., 2005; Duke et al., 2006; Elstner et al., 2009; Ohnuki et al., 2010; Gründemann et al., 2011; Botta-Orfila et al., 2012). A microarray study focusing on inflammatory gene expression in PD SN showed upregulation of the microglial purinergic receptor P2X7 (a receptor for ATP), colony stimulating factor-1 receptor (CSF1R), (a microglia growth factor receptor), and nitric oxide synthase 3 (a vascular marker) (Durrenberger et al., 2012).

With the development of high-sensitivity multiplex antibody arrays and other proteomic techniques, it is possible to profile large numbers of different biologically-active proteins in human brain tissue or other biological samples. We used this approach to examine the levels of 158 proteins in SN and striatum of control, ILBD and PD cases to determine if there were progressive changes in inflammation or related proteins. We particularly sought to determine if any of the cytokines identified in PD animal models could be validated in these human tissues. Our results demonstrated distinctly different patterns of inflammation and growth factor changes between SN and striatum with disease.

## MATERIALS AND METHODS

### Brain Tissue Samples

Brain tissue samples for this study were provided by the Banner Sun Health Research Institute Brain and Body Donation Program. The Brain and Body Donation Program operated with the approval of Western IRB (Puyallup, WA) under contract as the Institutional Review Board of Banner Research. A summary of the demographics of the cases is shown in **Table 1**. There were SN samples from 16 controls, 21 ILBD, and 18 PD cases; and striatum samples from 16 controls, 17 ILBD, and 16 PD cases. Tissue samples from both SN and striatum were not available for all cases—overlap of cases between brain regions was 92%. The selection of control, ILBD and PD cases used in this study was based on neuropathology diagnosis with reference to clinical records, with a diagnosis of Alzheimer's disease (AD) as the principal exclusion criteria. The degree of Lewy body (LB) pathology was assessed using a histological staging scheme in each of 10 brain regions (Beach et al., 2009). This involved obtaining a ranking score (0–4) using phosphorylated-alpha synuclein stained sections from each region. These numbers are summed to give Lewy body pathology scores of 0–40 for each brain. Histological ranking scores of plaques and tangles in five brain regions were used to assess how much age-associated AD-type pathology was present in each case (Beach et al., 2012; **Table 1**). This involved obtaining a ranking score (0–3) using Thioflavin S-stained tissue sections from entorhinal cortex, hippocampus, frontal, parietal, and temporal cortex. These numbers are summed to give plaque and tangle scores of 0–15 for each brain. Dementia was present in a number of the PD cases (**Table 1**); however, neuropathology and clinical records indicated this was not due to AD.

#### TABLE 1 | Demographic features of cases used in study.

*ILBD, incidental Lewy body disease; PD, Parkinson's disease; n, number of cases; M/F, male/female; LB, Lewy body staging score; Plaques, Plaque score; Tangles, tangle score; D, demented; MCI, mild cognitive impairment; CN, cognitively normal; DD, disease duration/years—mean and range.*

### Tissue Preparation

Due to the limited amount of SN tissue available from each case, SN tissue was provided as 15 consecutive frozen (10µm) SN sections (∼6–10 mg) cut from each block with a cryostat and collected frozen. This approach to tissue sample preparation allowed the selection of matched samples containing approximately equivalent densities of SN pars compacta neuromelanin-containing neurons. Frozen striatum samples (20–30 mg) were dissected from putamen at the level of the lenticular nucleus.

#### Processing of Tissue Samples

Both series of tissue samples were extracted in six volumes (w/v) of a proprietary extraction buffer compatible with the Quantibody arrays (Raybiotech, Norcross, GA) supplemented with protease/phosphatase inhibitors (Thermo-Fisher/Pierce). Samples were briefly sonicated in extraction buffer and incubated on ice with constant shaking for 30 min. After centrifugation (18,000 g/30 min), the supernatants were collected for further analysis. Protein concentrations were determined using a micro BCA assay (Thermo-Fisher/Pierce). These same extracts were also used to prepare western blot samples.

#### Tissue Analyses

Preliminary analyses of protein extracts were carried out using western blot methods for tyrosine hydroxylase (TH), as a measure of degree of dopaminergic neuron cell loss; for IBA-1, an indicator of microglia abundance; for TLR-2 as a marker for inflammation; and for glial fibrillary acidic protein (GFAP), as an indicator of reactive astrocytosis. We employed our standard published western blot methods using antibodies to TH (Biolegend, Dedham, MA; rabbit polyclonal 1:2000 dilution); to IBA-1 (Wako Chemicals, Richmond, VA, 1:2000); to TLR-2 (Abcam, Cambridge, MA; rabbit monoclonal 1:2500 dilution); and to GFAP [BD Biosciences, Franklin Lakes, NJ; cocktail of three different monoclonal antibodies (1:2000)] (Walker et al., 2015a,b).

For the antibody array analysis, samples were processed as a service by Raybiotech (Norcross, GA). For SN, each sample protein concentration was adjusted to 500µg/ml, and for striatum, each sample was adjusted to 1000µg/ml. These were analyzed using 160 protein Quantibody arrays (catalog number QAH-CAA-3000) composed of antibody-coated glass slide arrays for detecting cytokines, chemokines, soluble receptor, and growth factors. **Table 2** shows the arrangement of the proteins on the four separate array slides. Each protein had a standard curve included with measurements from known dilutions of purified standard protein. Each slide array contained positive control samples that were used for normalization purposes. Measurements were based on fluorescent intensity of bound labeled antibodies to each spot, and calculated from the mean of four spots/antibody. Slides were measured, analyzed, and normalized to positive controls using Raybiotech software. Final results were expressed as pg of protein/ml extract.

### Immunohistochemical Staining of SN and Striatum Tissue Sections

To characterize features of microglia activation in tissues being examined by antibody array analysis, 40µm paraformaldehydefixed tissue sections of SN and striatum from control, ILBD and PD cases were stained using antibodies LN3 (Abcam, Cambridge, MA) and IBA-1 (Wako Chemicals, Richmond, VA). These are well-established markers for activated microglia (LN3) or pan microglia (IBA-1). Striatum sections were double-stained for TH (Biolegend—rabbit polyclonal, 1:2000) using a two-color method. Immunohistochemstry was carried out according to our published procedures (Walker et al., 2009, 2015b).

### Data Analysis

The validity of the standard curve for each protein was confirmed by visual inspection of data. We found that standard curves for basic fibroblast growth factor and insulin growth factor binding protein-4 were unsatisfactory as the standard proteins did not produce a suitable dose-response curve; the results obtained for these proteins in tissue samples were excluded.

Data for each brain region samples were grouped as control, ILBD and PD and analyzed by One-way analysis of variance (ANOVA) using Graphpad Prism 5 software (Graphpad software, La Jolla, CA) without corrections for multiple comparisons. To

#### TABLE 2 | Quantibody array component proteins.

*Abbreviations/Definitions: Supplemental Table 1.*

determine whether age of patient or postmortem interval (PMI) were covariants for each measure, data from each protein was also analyzed by Analysis of Covariance (ANCOVA) with age, PMI, or age and PMI as covariants using Medcalc statistical software (Medcalc Software, Ostend, Belgium). Further analyses included correlation analyses, stepwise logistic regression analyses and receiver operating characteristic (ROC) curve analyses and were carried out using MedCalc. For each measure, P < 0.05 was considered statistically significant.

### RESULTS

### Characterization of Samples

The samples were selected based on consensus clinical and neuropathological criteria, but as shown in **Table 1**, there were different amounts of age-associated pathology in the samples. The control samples were free of LB pathology; while the ILBD and PD cases had varying degrees (**Table 1**). To support the clinical and neuropathological criteria used for case selection, samples were characterized for TH levels as an additional index of disease severity. There was significant variability between the samples in each disease group for TH, especially within the control groups (**Figure 1**). In SN, disease group differences in TH levels did not reach statistical significance by One-way ANOVA (**Figure 1A**), while in striatum, expected TH differences between each of the disease groups were shown (**Figure 1B**). Age and PMI were not significant covariant factors affecting TH levels in SN or striatum. Pathological variability within the disease groups was also highlighted by measures of gliosis and inflammation. GFAP levels showed no significant differences between disease groups for SN (**Figure 1C**) or striatum (**Figure 1D**) samples; however, GFAP levels in striatum were significantly affected by PMI (P = 0.03). There was significant negative correlation between TH and GFAP levels in striatum (Pearson r = −0.399, P = 0.0071; **Figure 1E**) suggesting increased gliosis as PD pathology progresses. Western blots measures of the microglial marker IBA-1 in SN (**Figure 1F**) and striatum (data not shown) did not show significant disease group differences; these measures were not affected by age or PMI.

#### Inflammatory Profiling of SN and Striatum

Initial data analysis of results from the Quantibody arrays consisted of individual ANOVA for each protein measure between the three disease groups for each of the brain regions. Results are shown in **Table 3**, which lists all P-values between groups for each protein. These results were not corrected for multiple comparisons. **Table 3** also identifies the proteins that gave negative values for each brain region. A negative designation was applied when fewer than 10 samples out of the total analyzed were above the limit of detection (LOD). For striatum samples, 57 of the 158 proteins measured were considered negative, while for SN, the same 57 were negative along with an additional 20 proteins (**Table 3**).

#### Protein Profile of SN

There were four proteins [IL-5, IL-15, monokine induced by gamma interferon (MIG) and IL-6 soluble receptor (IL6sR)] that gave P < 0.05 values between disease groups. The individual results for these proteins are shown as scatter plots in **Figure 2**. There were increases of IL-5, IL-15, and MIG, and a decrease of IL-6sR, in PD samples compared to controls or ILBD samples. Disease group differences did not become significantly different for any of these measures when corrected for age or PMI by ANCOVA; however, this analysis showed that IL-15 levels were significantly affected by PMI (ANCOVA—P = 0.040). In addition, when platelet derived growth factor (PDGF)-BB levels were corrected for age (P = 0.031—ANCOVA), a significant

disease group difference was obtained (P = 0.0323 between ILBD and PD samples).

Stepwise logistic regression analysis of these four cytokines was carried out to determine if these measures had predictive value between disease groups. Comparing control to PD indicated that only IL-5 values had significant predictive value (P = 0.003) with ROC curve analysis for IL-5 with sensitivity of 85.7% and specificity of 62.5% with area under curve (AUC) of 0.81. Further stepwise logistic regression analysis between ILBD and PD values showed a combination of IL-15 and IL-6sR levels gave predictive value (P = 0.0016), with ROC AUC of 0.820, and sensitivity of 85.7% and specificity of 64.7%. These stepwise regression analysis models were not strengthened by inclusion of TH values. Results are summarized in **Table 4**. Multiple stepwise regression analyses for all measured proteins in SN showed no additional proteins or group of proteins could discriminate between control and ILBD cases, or between ILBD and PD cases (data not shown).

#### Protein Profile of Striatum

A panel of 13 proteins was shown to have significant differences in striatum samples between the disease groups by Oneway ANOVA (**Table 3**)—CCL28 (C–C motif ligand 28 or mucosae-associated epithelial chemokine), HCC-1 (CCL-14), IL-18 binding protein a (IL-18BPa), PF4 (platelet factor 4), interferon-gamma (IFN-γ), IL-1 receptor antagonist (IL-1ra), IL-2, IL-15, PDGF-AA, PDGF-BB, TNF-α, Dtk (tyrosine-protein kinase receptor TYRO3), and ErbB3 (also HER3, human epidermal growth factor receptor 3). The individual scatter plot results for these proteins are shown in **Figure 3**. When correcting

#### TABLE 3 | Summary of One-way ANOVA results.