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The UPRmt Protects C. elegans From Mitochondrial Dysfunction by Upregulating 1

Specific Enzymes of the Mevalonate Pathway 2

Olga Oks1, Shany Lewin1, Irina Langier Goncalves, and Amir Sapir* 3

Department of Biology and the Environment, Faculty of Natural Sciences, University of 4

Haifa, Oranim, Tivon, 36006 Israel. 5

1These authors contributed equally to this work 6

* Corresponding author. E-mail: amirsapir1@gmail.com 7

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Running title: UPRmt upregulates mevalonate pathway enzymes 23

Genetics: Early Online, published on March 29, 2018 as 10.1534/genetics.118.300863

Copyright 2018.

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Keywords: Mitochondrial stress, UPRmt, Mevalonate pathway, Statins, Cholesterol. 1

Corresponding author: Amir Sapir, The Department of Biology and the Environment 2

University of Haifa, Oranim. Derch Kiryat Amal, Kiryat Tivon 3600600, Israel. 3

Phone: 972 495 396 15. E-mail: amirsapir1@gmail.com 4

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Abstract 23

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The mevalonate pathway is the primary target of the cholesterol-lowering drugs statins, 1

some of the most widely prescribed medicines of all time. The pathway’s enzymes not 2

only catalyze the synthesis of cholesterol but also of diverse metabolites such as 3

mitochondrial electron carriers and isoprenyls. Recently, it has been shown that one 4

type of mitochondrial stress response, the UPRmt, can protect yeast, C. elegans, and 5

cultured human cells from the deleterious effects of mevalonate pathway inhibition by 6

statins. The mechanistic basis for this protection, however, remains unknown. Using C. 7

elegans, we found that the UPRmt does not directly affect the levels of the statin target 8

HMG-CoA reductase, the rate-controlling enzyme of the mevalonate pathway in 9

mammals. Instead, in C. elegans the UPRmt upregulates the first dedicated enzyme of 10

the pathway, HMG-CoA synthase (HMGS-1). A targeted RNAi screen identified two 11

UPRmt transcription factors, ATFS-1 and DVE-1, as regulators of HMGS-1. A 12

comprehensive analysis of the pathway’s enzymes found that, in addition to HMGS-1, 13

the UPRmt upregulates enzymes involved with the biosynthesis of electron carriers and 14

geranylgeranylation intermediates. Geranylgeranylation, in turn, is requisite for the full 15

execution of the UPRmt response. Thus, the UPRmt acts in at least three coordinated, 16

compensatory arms to upregulate specific branches of the mevalonate pathway, 17

thereby alleviating mitochondrial stress. We propose that statin-mediated inhibition of 18

the mevalonate pathway blocks this compensatory system of the UPRmt and 19

consequentially impedes mitochondrial homeostasis. This effect is likely one of the 20

principal bases for the adverse side effects of statins. 21

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Introduction 1

In eukaryotes, the mevalonate/isoprenoid/cholesterol synthesis pathway, hereafter 2

referred to as the mevalonate pathway, catalyzes the synthesis of metabolites vital for 3

cellular metabolism, growth, and differentiation (SCHOENHEIMER AND BREUSCH 1933; 4

BLOCH 1965). The main branch of the pathway converts acetyl-CoA to farnesyl 5

diphosphate, which then serves as a precursor for the metabolism of most sub-6

branches. At each sub-branch, the sequential function of one of a series of metabolic 7

enzymes converts the farnesyl diphosphate precursor to a different end product. In 8

mammals, these end products include hemeA and ubiquinones, both of which act as 9

electron carriers for the mitochondrial electron transfer chain (ETC); dolichol, a lipid 10

anchor for N-glycosylation; isoprenyls, which are used for protein prenylation; and 11

cholesterol (reviewed in (GOLDSTEIN AND BROWN 1990)). Mevalonate pathway 12

metabolites are essential for many aspects of cellular and organismal metabolism, and 13

thus the function or malfunction of the pathway has fundamental implications for human 14

health and disease. For example, a loss in isoprenylation can promote apoptosis (ARAKI 15

et al. 2012), inhibit tumorigenesis through its effects on GTPases (JIANG et al. 2014), or 16

can lead to myopathy (DIRKS AND JONES 2006). Similarly, while cholesterol is essential 17

for plasma membrane integrity and fluidity, impaired cholesterol homeostasis promotes 18

the progression of many cardiovascular diseases (WOLLAM AND ANTEBI 2011). 19

More than four decades since their discovery (ENDO et al. 1976), statins remain a 20

leading clinical solution for patients with high serum cholesterol and are now among the 21

most widely used medications in the world. Statins are primarily used to lower the level 22

of cholesterol in the serum of patients through the inhibition of the second enzyme of 23

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the mevalonate pathway, HMG-CoA reductase. Statins, however, may have additional 1

beneficial effects, including having anti-cancer (CHAN et al. 2003; FREED-PASTOR et al. 2

2012) and anti-neurodegenerative properties (WANG et al. 2011). Although tens of 3

millions of people worldwide are currently prescribed with statins, an even more 4

widespread prescription of statins has been tempered because of their adverse side 5

effects. These effects range from mild muscle pain to severe pathological conditions 6

such as depression, insomnia, neuropathy, myopathy, and antiatherogenic syndrome 7

(GOLOMB AND EVANS 2008). A growing body of evidence suggests that the inhibition of 8

cholesterol synthesis itself may not be the primary cause of these deleterious side 9

effects. Instead, the inhibition of protein prenylation (WANG et al. 2008; CORREALE et al. 10

2014) and an impaired mitochondrial homeostasis (GOLOMB AND EVANS 2008; 11

APOSTOLOPOULOU et al. 2015) may be among the underlying mechanisms leading to the 12

side effects of statins. One proposed mechanism suggests that reduced Coenzyme Q10 13

(CoQ10) levels cause impaired mitochondrial calcium homeostasis in the muscles of 14

statin-treated humans (GALTIER et al. 2012; SIRVENT et al. 2012), but this is highly 15

controversial (APOSTOLOPOULOU et al. 2015; AUER et al. 2016). At present, the exact 16

underlying mechanism leading to the side effects of statins remains unknown. 17

Despite the mevalonate pathway’s centrality in human metabolism and its 18

reliance on the sequential activation of more than thirty enzymes, so far only a handful 19

of these enzymes were identified as being subject to regulation (SHARPE AND BROWN 20

2013). For example, milestone discoveries by Joseph L. Goldstein and Michael S. 21

Brown first demonstrated that the levels of cholesterol and cholesterol byproducts 22

regulate the metabolism of the pathway via negative feedback mechanisms (reviewed in 23

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(GOLDSTEIN AND BROWN 1990)). Paradigms describing the regulatory mechanisms of the 1

pathway typically focus on the transcriptional and post-translational regulation of the 2

second-most upstream enzyme of the pathway, HMG-CoA reductase (also known as 3

HMGCR1). Because HMGCR1 is the rate-determining enzyme of the pathway, its 4

regulation can determine the level of mevalonate pathway metabolism (REYNOLDS et al. 5

1984; GOLDSTEIN AND BROWN 1990; BURG AND ESPENSHADE 2011). In addition to 6

regulation through HMGCR1 activity, cholesterol and its derivatives affect the levels of 7

different enzymes of the cholesterol synthesis sub-branch, including DHCR7 (PRABHU et 8

al. 2016) and SQEL (HIDAKA et al. 1990; FORESTI et al. 2013; ZELCER et al. 2014). 9

Additionally, sterol-independent mechanisms of regulation exist (SHARPE AND BROWN 10

2013). These include, for example, the p53-dependent activation of enzymes in the 11

pathway during specific cancers (FREED-PASTOR et al. 2012) and the post-translational 12

regulation of the first enzyme of the pathway, HMGS-1, in C. elegans (SAPIR et al. 13

2014). The widespread and pleiotropic effects of the different metabolites of this 14

pathway on cellular functions have complicated attempts to piece apart their regulatory 15

mechanisms beyond the control of cholesterol synthesis. Interestingly, like many other 16

invertebrates, C. elegans lacks the cholesterol synthesis sub-branch of the pathway 17

(VINCI et al. 2008). The absence of the sub-branch for cholesterol synthesis could 18

facilitate the discovery of previously uncharacterized regulatory mechanisms typically 19

masked by this sub-branch. 20

Recently, a growing body of research has suggested an intimate crosstalk 21

between mitochondria and mevalonate pathway metabolism. Studies have found that a 22

key stress response pathway that safeguards the mitochondria, the mitochondrial 23

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Unfolded Protein Response (UPRmt), is capable of affecting mevalonate pathway 1

metabolism during stress. The activation of UPRmt via ethidium bromide (EtBr) 2

preconditioning protects yeast, C. elegans, and human cells in culture from the 3

inhibitory effects of statins (RAUTHAN et al. 2013). In addition, both mevalonate pathway 4

metabolism and its downstream sub-branch of protein prenylation are proposed to be 5

requisite for the activation of the UPRmt (RAUTHAN et al. 2013; LIU et al. 2014; RANJI et 6

al. 2014). Thus, the UPRmt may upregulate the mevalonate pathway, which in turn may 7

be essential for the proper full execution of the UPRmt. Nevertheless, the molecular 8

mechanisms underlying this proposed crosstalk between the UPRmt and the mevalonate 9

pathway remain unknown. 10

As one of the central metabolic centers of the cell, mitochondria are sensitive to 11

many external and internal insults, including energy crises (e.g. low levels of ATP) and 12

the accumulation of Reactive Oxygen Species (ROS) (HAYNES et al. 2013). The UPRmt 13

has evolved to protect the cell from these types of stressors (MELBER AND HAYNES 14

2018). In C. elegans, the UPRmt has been well characterized and many of its key 15

players are known (RUNKEL et al. 2013; MUNKACSY et al. 2016). In response to 16

mitochondrial or metabolic stress, the bZIP transcription factor ATFS-1 (HAYNES et al. 17

2010) translocates to the nucleus, where it induces a genome-wide transcriptional 18

response (NARGUND et al. 2012). ATFS-1 transcriptional targets include many 19

mitochondrial and cytoplasmic-localized chaperones, subunits of the electron transfer 20

chain, antioxidants, and mediators of innate immunity (PELLEGRINO et al. 2014). 21

Importantly, ATFS-1 activation alters the expression of enzymes in diverse metabolic 22

pathways (NARGUND et al. 2012), suggesting that it can remodel or even rewire 23

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metabolic networks. ATFS-1 works with a group of co-regulators in order to fine-tune 1

the UPRmt (HAYNES et al. 2013). Additional transcription factors that play a role in the 2

UPRmt response include the homeodomain protein DVE-1 (HAYNES et al. 2007), the 3

bZIP protein ZIP-2 (ESTES et al. 2010), and the bZIP protein SKN-1 (MUNKACSY et al. 4

2016). 5

In addition to the link between mevalonate pathway metabolism and the UPRmt, 6

inhibition of the mevalonate pathway by statins impairs farnesylation, thereby inducing 7

an ER stress response (MORCK et al. 2009). This activation of the ER stress-response 8

depends upon the activity of two canonical players in the UPRer, ire-1 and xbp-1 (Pilon 9

2009). Thus, statins activate the UPRer response while blocking UPRmt activation due to 10

a lack of geranylgeranylation. Nevertheless, the link between statins and the activation 11

of different stress signals deserves additional exploration. 12

In this work, we used C. elegans to understand better the relationship between 13

mitochondrial function and the mevalonate pathway. To identify the molecular 14

mechanisms that link mitochondrial dysfunction to mevalonate pathway metabolism, we 15

asked whether the mitochondrial stress response directly controls the steady-state 16

expression of enzymes in the mevalonate pathway. Intriguingly, we found that, in C. 17

elegans, mitochondrial dysfunction does not affect the levels of HMGR-1 protein, the 18

rate-controlling enzyme of the mevalonate pathway and the primary regulatory point in 19

mammals. Instead, the mitochondrial stress response upregulates the first dedicated 20

enzyme of the pathway, HMGS-1. We characterized the molecular basis for HMGS-1’s 21

upregulation and found that this effect relies upon ATFS-1 and DVE-1, two transcription 22

factors that mediate the UPRmt in C. elegans. Pathway-level transcriptional analyses 23

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identified at least three additional points of direct regulation of mevalonate pathway 1

metabolism by the UPRmt response. These include: i) upregulation of an enzyme of the 2

hemeA synthesis sub-branch; ii) upregulation of one enzyme, coq-1, that mediates the 3

first reaction in the sub-branch of ubiquinone production; and iii) upregulation of a 4

geranylgeranyl transferase. Finally, we show that deficiencies in the production of 5

ubiquinone and hemeA induce the UPRmt, which then upregulates mevalonate pathway 6

enzymes such as hmgs-1. This upregulation is dependent on atfs-1 activity that 7

functions in a compensatory circuit for the perceived mitochondrial dysfunction. 8

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Materials and methods 10

Worm strains and maintenance 11

Unless otherwise stated, C. elegans strains were maintained on NGM (Nematode 12

Growth Medium) plates at 20˚C, as previously reported (BRENNER 1974). The worm 13

strains used in this study and details of strain construction are provided in Table S1. 14

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Analysis of deletions and point mutations (dCAPS) 16

Deletions and point mutations were analyzed by the single-worm PCR method. A list of 17

primers is provided in Table S2. For the analysis of point mutations, the dCAPS method 18

was employed (NEFF et al. 1998) using the dCAPS Finder 2.0 software (NEFF et al. 19

2002). 20

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Pharmacological experiments 22

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All pharmacological experiments were conducted using 35mm diameter plates poured 1

with 2ml autoclaved NGM solution that was cooled to 55˚C and mixed with the different 2

pharmacological agents. For chemicals that were dissolved in solvents other than water, 3

the highest concentration of the solvent was used as a vehicle control. After drying the 4

plates for one day at room temperature, 150l of bacteria (OP-50 or other bacterial 5

strains) was seeded on the plate. After 24 hours at room temperature, the seeded 6

plates were used for experiments or stored in the refrigerator at 4˚C for a maximum of 7

two weeks. To initiate an experiment, approximately a hundred worms, synchronized to 8

larval stage one (L1) by incubation in a S. basal medium (STIERNAGLE 2006) for 24 9

hours post-bleach, were added to the plates. Unless stated otherwise, all experiments 10

were conducted at 20˚C and worms were analyzed 72-96 hours after the seeding of 11

worms on the plates, which corresponds to a chronological age of day 1 or day 2 of 12

adulthood. Tunicamycin (Sigma T7765) was dissolved in DMSO to a stock solution of 13

5mg/ml that was stored in -20˚C in the dark until used. The stock solution was diluted 14

1:1000 to a final concentration of 5g/ml in NGM plates and was compared to a control 15

NGM plates with DMSO diluted 1:1000. Mevalolactone (Sigma M4667) was dissolved in 16

water to create a stock solution of 1M and was used in final concentrations of 1, 10, or 17

20mM in the NGM plates. 18

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Feeding RNAi 20

All dsRNA bacterial clones were sequenced before use. Feeding was conducted as 21

previously described (TIMMONS et al. 2001) with the following modifications: i) Before 22

every experiment, we transformed HT115 (DE3) bacteria with the dsRNA expressing-23

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plasmids of interest. Freshly transformed bacteria (up to one week old) were picked and 1

cultured overnight in a water bath that was set to 37˚C in 2ml of Luria Broth (LB) 2

containing 50l/ml of ampicillin and 12.5l/ml of tetracycline. ii) The next morning, 3

bacterial cultures were diluted 1:100 in LB with 50l/ml ampicillin and were grown for six 4

hours in a 37˚C water bath. iii) IPTG (at a final concentration of 1mM) and ampicillin (at 5

a final concentration of 50l/ml) were spread on 35 or 60mm NGM plates before 6

seeding the bacteria. After six hours of incubation, a 1.5ml of bacterial culture per each 7

plate was harvested by spinning down at 4000 rpm. The bacterial pellet was 8

resuspended in 100l of LB and spread onto the plate. Plates were incubated for 12 9

hours in the dark at room temperature, and approximately 100 or 300 worms, 10

synchronized to the L1 stage by incubation in S. basal medium for 24 hours post-11

bleach, were added to the 35 or 60 mm RNAi plates respectively. In case of dsRNA 12

clones that affect worm survival (e.g. dsRNA corresponding to the hmgs-1 gene 13

sequence), the plates were seeded with 600 worms. Unless stated otherwise, all 14

experiments were conducted at 20˚C and were analyzed 72-96 hours after seeding the 15

worms in plates, when the worms were at the chronological age of day 1 or day 2 16

adults, respectively. For dsRNA mixing experiments, the bacterial clones were grown 17

separately and mixed in 1:1 (volume:volume) before seeding. 18

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Expression analysis by qPCR 20

Worms were bleached and synchronized by incubation for 24 hours in S. basal medium 21

at 20˚C. Synchronized wild-type worms at the L1 stage (approximately 300 worms/plate, 22

four plates per condition) were spotted onto empty vector and spg-7 RNAi plates. 23

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Worms were grown until day 1 of adulthood, then collected and washed with a S. basal 1

medium three times and with RNase-free water two more times. Total RNA was 2

extracted using a combination of Trizol/chloroform and an RNeasy kit (Qiagen) and was 3

treated with DNase I (New England Biolabs) according to the manufacturer instructions. 4

The concentration of the mRNA was quantified by both Nanodrop and Qubit (Thermo-5

Fisher Scientific), and 0.6μg of total RNA was used for cDNA synthesis using the 6

ProtoScript® II Reverse Transcriptase kit (New England Biolabs) with random hexamers 7

as the cDNA primers. qPCR reactions were performed using the Fast SYBR Green 8

Mastermix (ABI) in Step One Plus (ABI), or CFX Connects (BioRad) machines. Primers 9

were designed using the NCBI primer design tool (Primer-Blast) (YE et al. 2012). All 10

primers span at least one intron-exon junction and have a melting temperature (TM) 11

ranging from 58˚C to 62˚C. All primers were synthesized by IDT (Germany) and can be 12

found along with their calculated TM and amplicon size in Table S2. Primer sequences 13

for act-1 or pmp-3 were used for expression normalization (HOOGEWIJS et al. 2008; 14

ZHANG et al. 2012). The ΔΔCt method was used to calculate the fold change in 15

expression. Results are averages of three biological replicates, and error bars represent 16

standard errors of the mean. Statistical significances were calculated using the Prism 17

software with unpaired two-tailed Student’s T-tests. 18

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Fluorescence imaging and immunoblotting 20

To determine the level of expression of GFP, synchronized worms harboring different 21

GFP reporters were mounted on agar pads (2% agar in S. basal) with 3l of 10mM 22

levamisole (Santa Cruz, 16595-80-5). Unless otherwise stated, gravid adult worms were 23

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visualized at a chronological age of day one or two of adulthood, i.e., 72 and 96 hours 1

post-release from L1 arrest, respectively. For each condition, at least 30 worms were 2

analyzed, and at least ten worms were imaged in a group across all independent 3

experiments. For imaging of groups of worms on slides, ten worms were collected from 4

the plates and transferred to a 1l drop of the levamisole solution. Once the drop was 5

dried out, the worms were oriented head-to-head using a pick and then covered by a 6

coverslip. Levamisole solution was added from the margins of the slide until the entire 7

agar pad was covered with this solution, and the slides were kept for about 30 minutes 8

in a humid chamber to reduce the number and size of air bubbles. In every experiment, 9

the same magnification, light intensity, and exposure time were used. Images were 10

captured using a Nikon DSRi-1 camera connected to a Nikon E600 fluorescence 11

compound microscope or using a Nikon SMZ18 fluorescence-dissecting microscope 12

connected to a Nikon DS-Fi3 camera. Images were processed using the Adobe 13

Photoshop software according to ethical guidelines for the appropriate use and 14

manipulation of scientific digital images (CROMEY 2010). 15

For Immunoblotting, worms were grown on spg-7 or empty vector RNAi plates. At 16

the adult stage, 50 worms were collected, lysed, and the total proteome was separated 17

by SDS-PAGE electrophoresis. Next, the proteins were transferred to nitrocellulose 18

membranes, and probed for the HMGS-1::GFP protein using an anti-GFP antibody 19

followed by stripping and probing with an anti-actin antibody. 20

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Determination of the number of nuclei with DVE-1::GFP 22

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Worms expressing the DVE-1::GFP construct were mounted as described above. 1

Nuclei with GFP signal were counted in a Nikon Eclipse 600 microscope using a 100X 2

magnification. The statistical significance of the change in the number of DVE-1::GFP 3

nuclei in each condition was calculated by one-way ANOVA testing using the SPSS 4

software. Normality was tested and confirmed by the Shapiro-Wilk test. Next, the 5

Scheffe post-hoc test was included to rule out the effect of multiple tests. 6

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Thrashing assay 8

Worms until a chronological age of day one adult were grown as described above. 9

Groups of five worms were placed in a drop of 20l S. basal medium on a glass slide, 10

and after about five minutes of habituation, the number of thrashes was measured. 11

Movies of the swimming worms, in thirty seconds intervals, were captured using Nikon’s 12

SMZ18 dissecting microscope and the Active Presenter software. The captured movies 13

were then played in slow motion for 20 seconds to count the number of times the head 14

or the tail cross the midline of the worm, which we defined as one thrash. The statistical 15

significance of the results was analyzed by one-way ANOVA testing using SPSS 16

software. Normality was tested by the Shapiro-Wilk test followed by the Kruskal-Wallis 17

test to analyze data that are not normally distributed. 18

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Generation of a deletion allele in the hmgs-1 locus using the CRISPR-Cas9 20

technology and rescue experiments 21

Deletion was generated, upon our request, in the laboratory of Donald G. Moerman as 22

part of the CRISPR-Cas9 Gene Knockout project in C. elegans. An integration cassette, 23

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including homology arms in the hmgs-1 gene and a gfp gene under the regulation of a 1

myo-2 promoter was used. This cassette replaced the sequence of the hmgs-1 gene 2

from the middle of exon two through the middle of exon seven. This replacement 3

removed the majority of the eight total exons of the hmgs-1 gene introducing a 4

frameshift after Serine 64 and a stop codon ten further amino acids downstream. The 5

replacement was validated by PCR and worms having this deletion allele were 6

outcrossed four more times to N2 wild-type worms before the experiments. 7

Supplementation of 20mM mevalolactone to the NGM plates can fully rescue the 8

lethality associated with this hmgs-1 deletion allele providing another indication for the 9

specificity of this allele. To test the recuse potential of the hmgs-1::gfp transgene, this 10

integrated construct was recombined with the hmgs-1 deletion allele. Worms 11

homozygous for the hmgs-1 deletion allele, either with or without the hmgs-1::gfp 12

construct, were grown on plates with 20mM mevalolactone. These worms, at larval 13

stage four, were transferred to plates without mevalolactone and the number of progeny 14

and livelihood of the worms were determined 96 hours after the transfer. 15

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Data Availability Statement 17

Strains and plasmids are available upon request. Table S1 contains data about all the 18

C. elegans strains used in this study. Table S2 contains data and the individual 19

sequences of all the primers used in this study. The results of the screen conducted to 20

find GTPases that regulate the levels of HMGS-1::GFP are presented in Table S3. 21

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Results 23

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Mitochondrial Stress Upregulates hmgs-1 1

To determine whether the mitochondrial stress response affects the metabolism of the 2

mevalonate pathway, we first measured the levels of different mevalonate enzymes in 3

vivo in C. elegans. We started with the first dedicated enzyme in the pathway, HMG-4

CoA synthase (HMGS-1 in C. elegans or HMGCS1 in mammals). This enzyme 5

undergoes transcriptional regulation in mammals (GIL et al. 1986) and an aging-6

dependent SUMOylation in C. elegans (SAPIR et al. 2014). First, we measured the levels 7

of HMGS-1 protein by using an integrated fosmid in which a green fluorescence protein 8

(gfp) coding sequence was recombined in-frame at the 3’ end of the hmgs-1 gene 9

(SAROV et al. 2012; SAPIR et al. 2014). In unstressed conditions, worms carrying this 10

construct (hereafter, HMGS-1::GFP) had a weak GFP signal in the intestine and even a 11

weaker signal in other tissues ((SAPIR et al. 2014); Fig. 1A). In contrast, after using RNAi 12

against the mitochondrial metalloprotease spg-7 (paraplegin) to induce mitochondrial 13

stress (NARGUND et al. 2012; PELLEGRINO et al. 2014), the HMGS-1::GFP signal was 14

strongly upregulated in the intestine (Fig. 1B). A similar upregulation of HMGS-1::GFP 15

(Fig. 1C, D) was seen in a strain harboring a mutation in the iron/sulfur protein isp-1 that 16

also results in mitochondrial stress (NARGUND et al. 2012; MUNKACSY et al. 2016). Under 17

each of these conditions, the mitochondrial stress response upregulated HMGS-1::GFP 18

primarily in the intestine. Double-stranded RNA corresponding to the hmgs-1 gene 19

eliminated the GFP signal in HMGS-1::GFP worms treated with spg-7 RNAi (Fig. S1A, 20

B), demonstrating the specificity of the HMGS-1::GFP construct and the RNAi 21

treatment. 22

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A growing body of evidence suggests that mitochondrial stress responses share 1

some molecular and cellular characteristics with other types of organelle stress 2

responses, such as those of the endoplasmic reticulum (ER) (HAYNES et al. 2013; 3

RUNKEL et al. 2013). Therefore, it is possible that other forms of stress such as the ER 4

stress response affect the levels of hmgs-1. To address these possibilities, we exposed 5

animals to a wide variety of stress-inducing conditions, ranging from starvation to ER 6

stress (UPRer). Importantly, we found no change in the level of HMGS-1::GFP across 7

any of the other conditions tested. For example, Tunicamycin-induced UPRer did not 8

alter HMGS-1::GFP distribution (Fig. S1C, D), whereas the same treatment induces the 9

upregulation of a reporter for UPRer (Fig. S1E, F). This strongly suggests that the 10

upregulation of hmgs-1 is highly specific to mitochondrial stress. 11

We next aimed to test whether the HMGS-1::GFP protein is active in vivo. To 12

assess this, CRISPR-Cas9 technology was used to generate a loss of function allele in 13

the hmgs-1 gene. CAS-9 activity induces the insertion of a cassette replacing the 14

sequence of hmgs-1 gene from the middle of exon two through the middle of exon 15

seven. This replacement resulted in the removal of the majority of the eight total exons 16

of the hmgs-1 gene (Fig. S2A-C) introducing a frameshift after Serine 64 in the HMGS-1 17

protein and a stop codon ten further amino acids downstream. Consistent with HMGS-1 18

having an essential function in the mevalonate pathway that supports organism 19

metabolism, growth, and development, animals homozygous for the hmgs-1 deletion 20

allele died as embryos (Fig. S2D). This lethality, along with the loss of most of the exons 21

of the hmgs-1 gene including exons that encode for the active site of the enzyme, 22

suggest that this hmgs-1 knockout is a complete loss-of-function allele. To determine 23

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the functionality of the HMGS-1::GFP construct in vivo, we introduced this construct in 1

the background of the hmgs-1 knockout allele (Fig. S2B-E). Importantly, the HMGS-2

1::GFP construct can rescue the lethal phenotypes of the hmgs-1 knockout allele, 3

resulting in the generation of worms that were successfully able to reach the adult stage 4

(Fig. S2D, E). This rescue demonstrates the activity of the HMGS-1::GFP protein in vivo 5

and suggests that the expression and distribution of this HMGS-1::GFP protein is similar 6

to the endogenous HMGS-1 enzyme. 7

Regulation of the mevalonate pathway in mammals relies primarily on the control 8

of the second enzyme of the pathway HMG-CoA reductase, or HMGCR1. Thus, we next 9

studied the putative ortholog of HMGCR1 in C. elegans, HMGR-1, by expressing a 10

HMGR-1::GFP fusion protein under the control of the hmgr-1 promoter. Consistent with 11

a previous report (RANJI et al. 2014), HMGR-1 protein was localized in several tissues, 12

including the worm intestine and spermatheca (Fig. 1E). Surprisingly, however, we 13

found that spg-7 RNAi was incapable of altering HMGR-1 expression levels or spatial 14

distribution (Fig. 1F). This was in stark contrast to the strong upregulation of HMGS-1 15

during mitochondrial stress (Fig. 1B, D). 16

HMGS-1 upregulation could stem from several mechanisms, including the 17

induction of transcription or an inhibition in degradation of the protein. To determine 18

whether an increase in the level of hmgs-1 transcription leads to the observed 19

upregulation of the HMGS-1::GFP protein, we analyzed the promoter activity of hmgs-1 20

in vivo by expressing the hmgs-1 promoter directly fused to the sequence of the gfp 21

gene. Upon mitochondrial stress, we detected an upregulation of the GFP signal in the 22

intestine of worms harboring this reporter (Fig. 1G, H). Although we cannot rule out 23

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some contribution by post-translational modifications, the strong correlation both 1

temporally and spatially between the promoter activity of hmgs-1 and HMGS-1::GFP 2

protein distribution suggests that transcriptional mechanisms account for HMGS-1 3

upregulation. We then used quantitative PCR (qPCR) to follow the levels of the 4

endogenous transcripts of hmgs-1 and hmgr-1 genes in wild-type worms stressed by 5

the spg-7 dsRNA in comparison to the empty vector control. In agreement with the 6

results of the GFP fusion constructs, we found that hmgs-1 mRNA was upregulated by 7

2.4-fold in spg-7 RNAi in comparison to the empty vector control (Fig. 1I). In contrast, 8

and in accordance with our analysis of the fusion protein in Fig. 1F, using qPCR we 9

observed no significant change in the level of hmgr-1 mRNA (Fig. 1I). The increase in 10

HMGS-1::GFP protein was confirmed using a western blot analysis (Fig. 1J) with a third 11

type of mitochondrial stress, RNAi against the ATP synthase gene atp-3 (VENTURA et al. 12

2009). 13

Finally, because HMGS-1 protein has not been well characterized in C. elegans, 14

we examined whether this protein is indeed a functional ortholog of the mammalian 15

HMGCS1 enzyme. HMGCS1 is capable of catalyzing the first dedicated biochemical 16

reaction of the mevalonate pathway—i.e. the conversions of Acetyl-CoA and 17

Acetoacetyl-CoA into HMG-CoA (SKAFF AND MIZIORKO 2010). To this end, we measured 18

the activity of the C. elegans HMGS-1 enzyme in vitro. We expressed and purified 19

HMGS-1 and human HMGCS1 proteins from bacteria (Fig. S3A) and found that both 20

proteins were capable of converting AcetylCoA and Acetoacetyl-CoA into HMG-CoA 21

(Fig. S3B). We also found that the treatment of C. elegans animals with mevalonate, a 22

metabolite synthesized downstream to HMGS-1, fully rescued the severe phenotypes 23

20

caused by hmgs-1 RNAi (Fig. S4A-F). The full rescue of hmgs-1 RNAi phenotypes by 1

mevalonate suggests that, in C. elegans, HMGS-1 acts solely in the mevalonate 2

pathway. Taken together, our in vitro and in vivo data indicate that HMGS-1 is a 3

functional ortholog of human HMGCS1, acting as the first dedicated enzyme of the 4

mevalonate pathway. Our results demonstrate that mitochondrial stress upregulates 5

hmgs-1 transcription and consequentially HMGS-1 protein levels as a mitochondrial 6

stress-specific response. Based on these results, we hypothesized that in the intestine, 7

the primary metabolic tissue of the animal, the mitochondrial stress response 8

upregulates HMSG-1 as part of a compensatory mechanism to restore homeostasis. 9

10

The UPRmt Transcription Factors ATFS-1 and DVE-1 Upregulate hmgs-1 11

The genetic amenability of C. elegans has led to the identification of a complex network 12

of transcription factors and co-regulators that form the basis of the mitochondrial stress 13

response (Fig. 2A). These include the transcription factors ATFS-1 (NARGUND et al. 14

2012), DVE-1 (HAYNES et al. 2007), SKN-1 (MUNKACSY et al. 2016), and ZIP-2 15

(PELLEGRINO et al. 2014). Additional proteins that play a major role in the mitochondrial 16

stress response are the transporter HAF-1 and the eIF2 kinase GCN-2, the latter of 17

which regulates protein translation during mitochondrial stress (BAKER et al. 2012). 18

To understand how mitochondrial stress response governs hmgs-1 19

upregulation, we conducted a targeted genetic screen to identify genes that could affect 20

hmgs-1 expression. First, we used an isp-1(qm150) mutant background to elicit 21

mitochondrial stress and to constitutively upregulate the HMGS-1::GFP protein (Fig. 22

2B). We then systematically tested 36 genes previously characterized as having a role 23

21

in the mitochondrial stress response (MUNKACSY et al. 2016; TIAN et al. 2016) for their 1

effects on hmgs-1 expression. We compared the HMGS-1::GFP signal of worms treated 2

with each dsRNA clone with the signal of the empty vector (Fig. 2B) or hmgs-1 RNAi 3

(Fig. 2C) controls. In these analyses, we found that three dsRNA clones, corresponding 4

to the sequences of the UPRmt related genes atfs-1, dve-1, and rheb-1, attenuated the 5

upregulation of HMGS-1::GFP in isp-1 mutant worms (Fig. 2D-F). 6

Previous studies have suggested that zip-2 (PELLEGRINO et al. 2014) and skn-1 7

(MUNKACSY et al. 2016) may act downstream of atfs-1 in the mediation of some of its 8

responses. Our genetic analyses, however, suggested that these genes were 9

dispensable for the isp-1(qm150) dependent upregulation of HMGS-1::GFP expression 10

(Fig. S5A-C). Similarly, the knockdown of gcn-2, which plays a role independent of atfs-11

1 in the mitochondrial stress response, did not affect the upregulation of HMGS-1::GFP 12

in the isp-1 mutant background (Fig. S5D). 13

Additional characterized effectors of mitochondrial or metabolic stress including 14

the daf-16 gene, the dlk-1/pmk-3/sek-3 pathway, and components of the mTOR 15

pathway also did not affect HMGS-1::GFP upregulation (Fig. 2G). Moreover, we did not 16

detect a change in HMGS-1::GFP levels when knocking down cep-1, the ortholog of 17

p53 that upregulates HMGCS1 in specific cancers (FREED-PASTOR et al. 2012), or sbr-1 18

the ortholog of the SREBP proteins that mediate the cholesterol-dependent regulation of 19

HMGCS1 (HORTON et al. 2002). Because the nature and dynamics of the UPRmt are 20

somewhat affected by the type of stressor, we also used the spg-7 RNAi as a second, 21

independent method of inducing the UPRmt (Fig. S6A-C). Consistent with the approach 22

of using the isp-1 mutation, we found that both atfs-1 (Fig. S6D) and dve-1 (Fig. S6E) 23

22

were required for the upregulation of the HMGS-1::GFP protein. All other components of 1

the UPRmt (Fig. S6F-J) tested by the RNAi approach were dispensable for the 2

upregulation hmgs-1 expression by spg-7 RNAi. Nevertheless, because RNAi typically 3

only partially reduces the expression of the corresponding gene, we cannot completely 4

rule out the possibility that genes found not to alter the levels of the HMGS-1::GFP 5

protein still have a minor role in HMGS-1 regulation. 6

These results strongly suggest that the molecular mechanisms of HMGS-1 7

upregulation are extremely context-dependent and highly regulated. Collectively, our 8

screen identified a specific regulation of HMGS-1 by UPRmt master regulators ATFS-1 9

and DVE-1 during mitochondrial stress. 10

11

Impaired HemeA and Ubiquinone Synthesis Results in UPRmt Activation and a 12

Compensatory HMGS-1 upregulation response 13

Why would HMGS-1 be upregulated during mitochondrial stress? We hypothesized that 14

this upregulation by the UPRmt may serve as a compensatory response, allowing the 15

cell to increase the synthesis of metabolites required to restore mitochondrial 16

homeostasis. To test this hypothesis, we knocked down different branches of the 17

mevalonate pathway while monitoring the level of UPRmt induction and HMGS-1::GFP 18

upregulation. We used the promoters of the mitochondrial chaperones hsp-6 and hsp-19

60 fused to the gfp gene (hsp-6pr::gfp and hsp-60pr::gfp respectively) to assess the level 20

of UPRmt induction (YONEDA et al. 2004). Consistent with previous reports (RAUTHAN et 21

al. 2013; LIU et al. 2014; RANJI et al. 2014), knocking down hmgs-1 or other upstream 22

enzymes of the main branch of the mevalonate pathway did not lead to the induction of 23

23

UPRmt (Fig. S7A). Similarly, RNAi against enzymes of the sub-branches that account for 1

tRNA modification, protein prenylation, or dolichol synthesis—although often resulting in 2

potent visible phenotypes—did not result in an induction of the UPRmt or HMGS-1 3

upregulation (Fig. S7A, B). In stark contrast, however, we found that knocking down an 4

enzyme in the sub-branch that facilitates hemeA biosynthesis (cox-15) triggered a 5

potent UPRmt induction (Fig. 3A, B), which was dependent on atfs-1 activity (Fig. 3C, D). 6

Because both hemeA and ubiquinone serve as carriers of electrons in the 7

electron transport chain (ETC), their loss may be indicative of a shared response. To 8

test this, we used loss-of-function mutations of coq-1 and coq-2, two genes encoding 9

enzymes of the ubiquinone synthesis pathway, to block ubiquinone biosynthesis. We 10

found that, while worms homozygous for coq-1 or coq-2 loss-of-function mutation 11

arrested at the young adult stage, they exhibited a strong induction of the hsp-6pr::gfp 12

reporter (Fig. 3E, F; Fig. S8A, B). Interestingly, in coq-1 mutant animals, hsp-6pr::gfp 13

upregulation was dependent on atfs-1 (Fig. 3G) and dve-1 (Fig. 3H), whereas in the 14

coq-2 mutant background this upregulation was dependent on atfs-1 (Fig. S8C) but only 15

marginally on dve-1 (Fig. S8D). The effect of coq-2 was independent of the bacterial 16

strains used as a food source and occurred equally in HT115 and OP50 strains (Fig. 17

S9A-D). 18

If a feedback loop mechanism exists, a reduction in the level of specific 19

metabolites of the pathway will activate a system that will upregulate enzymes 20

responsible for the synthesis of the deficient metabolites. To examine this possibility, we 21

analyzed the levels of hmgs-1 during reduced ubiquinone production by measuring the 22

levels of HMGS-1::GFP in coq-2 loss-of-function mutant animals. In consistence with 23

24

this model, we found that the loss of a functional ubiquinone synthesis pathway leads to 1

HMGS-1::GFP upregulation (Fig. S10A, B) and that this upregulation is partly 2

dependent on atfs-1 (Fig. S10C). Although ATFS-1 has many different targets 3

(NARGUND et al. 2012), the upregulation of HMGS-1::GFP has a mechanistic rationale 4

as a compensatory response in situations in which the production of ubiquinone and 5

hemeA is attenuated. Taken together, our results support a model of a compensatory 6

mechanism that has evolved to cope with low levels of ubiquinone or hemeA electron 7

carriers by inducing the UPRmt response to upregulate the HMGS-1 enzyme. 8

To understand the physiological significance of UPRmt activation following the 9

block in ubiquinone synthesis, we measured worm thrashing as a functional readout for 10

animal health. We found that a lack of ubiquinone synthesis significantly reduced the 11

number of thrashes, presumably because of a reduction in respiration or in other 12

mitochondrial functions that impede normal muscle activity. Although atfs-1 RNAi was 13

efficient in blocking the UPRmt signal in the coq-1 mutant background (Fig. 3G), it did not 14

significantly affect the level of worm thrashing (Fig. 3I). This further supports a model in 15

which the UPRmt directly upregulates enzymes of the mevalonate pathway. Because the 16

coq-1 allele used is a presumable null allele (RODRIGUEZ-AGUILERA et al. 2005), any 17

upregulation of enzymes, either up or downstream of coq-1, would not be able to rescue 18

the phenotypes of coq-1 deficiency. 19

20

UPRmt Requires the Metabolism of the Mevalonate Pathway Main Branch and 21

Geranylgeranylation 22

25

The above experiments raise a conundrum: the deficiency in hemeA and ubiquinone 1

production strongly elicits the UPRmt (Fig. 3B, F), but the inactivation of the main branch, 2

which presumably also results in hemeA and ubiquinone deficiency, fails to induce the 3

UPRmt (Fig. S7A). It is possible, therefore, that some products of the mevalonate 4

pathway are required for the UPRmt. This hypothesis is in agreement with recent 5

literature suggesting the necessity of a functional mevalonate pathway for UPRmt 6

activation (RAUTHAN et al. 2013; LIU et al. 2014; RANJI et al. 2014). 7

To test this hypothesis, we knocked down different branches of the pathway in 8

the background of a constitutively activated UPRmt response. We used the isp-1(qm150) 9

mutation to induce UPRmt and the hsp-6pr::gfp reporter as the readout for the level of the 10

UPRmt signaling (Fig. 4A). As a positive control, we used atfs-1 and dve-1 RNAis to 11

reduce the expression of the UPRmt reporter, as previously established (Fig. 4B, C). 12

Next, we tested whether reduced expression of enzymes in the main branch of the 13

pathway could affect the UPRmt. We found that knocking down hmgs-1 or hmgr-1 14

attenuated the activation of the UPRmt response in the isp-1 mutant background (Fig. 15

4D, E). In contrast, hmgs-1 RNAi did not attenuate the ER stress response (Fig. S11A-16

C), suggesting that the requirement of a functional mevalonate pathway is specific for 17

the UPRmt. Importantly, we were able to chemically rescue, in a concentration-18

dependent manner, the block in UPRmt activation caused by hmgs-1 RNAi by 19

supplementing mevalonate exogenously (Fig. S12A-F). This rescue directly suggests 20

that one of the metabolites produced by the pathway downstream of mevalonate itself is 21

required for UPRmt activation. 22

26

Next, we tested whether reduced expression of enzymes in any of the other sub-1

branches of the mevalonate pathway was sufficient to inhibit UPRmt induction. We found 2

that knocking down the ggtb-1 gene, which encodes a type II geranylgeranyltransferase, 3

inhibits UPRmt induction (Fig. 4F). In contrast, knocking down other sub-branches of the 4

mevalonate pathway did not attenuate the activation of the UPRmt response (Fig. S13). 5

The UPRmt relies on the sequential activation of several proteins, including ATFS-6

1, which plays a cardinal role in the UPRmt response. Therefore, impaired mevalonate 7

pathway metabolism concomitant with a block of geranylgeranylation may impede 8

ATFS-1 activation. To test the possible connection between mevalonate pathway 9

metabolism and ATFS-1 activation, we used a strain harboring a constitutively activated 10

allele of atfs-1 (atfs-1(et15)). In this mutant background, ATFS-1 protein translocates to 11

the nucleus, where it exerts its activity constitutively even in the absence of 12

mitochondrial stress (RAUTHAN et al. 2013; PELLEGRINO et al. 2014) (Fig. 4G). As a 13

control, we used the atfs-1 RNAi that blocked the UPRmt signal almost completely in 14

most of the tissues of these animals (Fig. 4H). We found that, in this background, 15

knockdown of hmgs-1 or ggtb-1 did not affect the level of UPRmt induction (Fig.4 I, J), 16

indicating that the requirement for mevalonate pathway metabolism and 17

geranylgeranylation lies upstream or in parallel to ATFS-1 activation. 18

Taken together, our results suggest that the reliance of the UPRmt response on a 19

functional mevalonate pathway metabolism stems from the requirement for functional 20

geranylgeranylation. Furthermore, our genetic analyses reveal that geranylgeranylation 21

is required upstream or in parallel to ATFS-1 activation. 22

23

27

Inhibition of the UPRmt Does Not Stem from RHEB-1 Inactivation 1

Geranylgeranylation is requisite for the activation of many small GTPases (CHERFILS 2

AND ZEGHOUF 2013), underscoring the possibility of the involvement of a small GTPase 3

in the activation of the UPRmt response. A growing body of evidence suggests an 4

evolutionary conserved role for GTPases as facilitators of many stress responses that 5

coordinate cellular metabolism (BAR et al. 2016). It is plausible, therefore, that in C. 6

elegans, the activation of the UPRmt relies on the function of a specific GTPase. Of note, 7

in our targeted screen, apart from ATFS-1 and DVE-1, the only other gene whose RNAi 8

altered the levels of HMGS-1 is the small GTPase rheb-1 (Fig 2F), which was previously 9

identified as regulator of the UPRmt (HAYNES et al. 2007). Based on these two indicators, 10

it is possible that impaired mevalonate pathway metabolism and loss of 11

geranylgeranylation block RHEB-1 activation and consequentially the UPRmt response. 12

To test this model, we examined whether mevalonate pathway metabolism is 13

requisite for RHEB-1 activation in the context of UPRmt. Similar to the hmgs-1 and ggtb-14

1 knockdown results, we found that rheb-1 knockdown somewhat modulates UPRmt 15

activation (Fig. S14 A, B). This effect of rheb-1 takes place upstream or in parallel to 16

atfs-1 activation (Fig. S14C-F). Next, we used the DVE-1::GFP reporter, previously 17

shown to be a readout for RHEB-1 protein function (HAYNES et al. 2007), to determine 18

the level of RHEB-1 activation. In wild-type worms, DVE-1::GFP accumulates for 19

unclear reasons at a few nuclei at the anterior and posterior sides of the worm (Fig. 20

S15A, A’). As previously reported (HAYNES et al. 2007), we found that RHEB-1 loss 21

leads to an increased number of DVE-1::GFP positive nuclei at the posterior side of the 22

worms (Fig. S15B, B’). The knockdowns of hmgs-1 and ggbt-1 result in severe 23

28

phenotypes that demonstrate the potency of these RNAis, but these conditions did not 1

alter the number of DVE-1::GFP nuclei in comparison to the control EV RNAi (Fig. 2

S15A, C-E). Although it is possible that the distribution of DVE-1::GFP reporter 3

represents only one aspect of RHEB-1 activation, this result suggests that impaired 4

prenylation does not affect at least one function of RHEB-1, i.e. restricting DVE-1::GFP 5

expression. 6

Because RHEB-1 did not appear to be the missing link between 7

geranylgeranylation and the UPRmt, we screened the family of small GTPases in C. 8

elegans for suppressors of the UPRmt response. In the background of a constitutively-9

activated UPRmt, i.e. the isp-1(qm150), hsp-6pr::gfp background, we tested 49 out of the 10

53 predicted small GTPases encoded by the C. elegans genome (REINER AND 11

LUNDQUIST 2016). To rule out a possible effect of the method used, we used an 12

additional approach to induce the UPRmt, i.e. the spg-7 RNAi. Using these two 13

approaches, we did not identify in our screen any gene, other than rheb-1, as being a 14

requirement for UPRmt activation (Table S3). 15

16

Mitochondrial Stress Induces a Coordinated Upregulation of Three Different Sub-17

branches 18

Our pathway-level genetic analyses demonstrated that proper mitochondrial function 19

requires hemeA and ubiquinone biosynthesis. Moreover, we found that 20

geranylgeranylation plays a critical role in the execution of the UPRmt response. These 21

results highlight the possibility that, along with HMGS-1 of the main branch, UPRmt 22

upregulates the levels of additional enzymes in order to increase the metabolic flux of 23

29

selected sub-branches. To address this possibility, we first examined available whole-1

genome expression data related to the UPRmt (NARGUND et al. 2012). Highly consistent 2

with our genetic analyses, these expression studies identified enzymes of the hemeA, 3

ubiquinone, and geranylgeranylation sub-branches as enzymes that are potentially 4

upregulated upon mitochondrial stress (Fig. 5A). To examine this possibility, at a single 5

gene resolution, we measured the levels of endogenous transcripts of different enzymes 6

of the pathway, comparing N2 wild-type worms fed with either the spg-7 dsRNA or an 7

empty vector control. In agreement with both our genetic analysis and the whole 8

genome studies, we found that mitochondrial stress specifically upregulates enzymes of 9

the three sub-branches (Fig. 5A, S16). These include an enzyme that catalyzes the 10

biosynthesis of a metabolite essential for the hemeA biosynthesis (Y46G5A.2), the first 11

enzyme of the ubiquinone synthesis sub-branch (coq-1), and one enzyme that mediates 12

the geranylgeranylation of proteins (M57.2). Next, we analyzed the expression and 13

distribution of selected proteins encoded by the upregulated genes in vivo during 14

mitochondrial stress. All of the upregulated genes we identified, except hmgs-1, lie 15

within operons and therefore cannot be fused to a gfp sequence using standard cloning 16

or fusion PCR methods. Nevertheless, one of these genes, coq-1, is available with a gfp 17

sequence at its 3’ end in the TransgeneOme library of fosmids (SAROV et al. 2012). 18

Therefore, we obtained and validated the integrity of the gfp-tagged coq-1 gene in the 19

fosmid. Next, we generated transgenic worms harboring the coq-1::gfp fosmid. COQ-20

1::GFP expressing worms exhibited a weak, intestinally-enriched GFP signal (Fig. 5B) 21

that is abolished by coq-1 RNAi (Fig. S17A, B). Consistent with our qPCR results, the 22

COQ-1::GFP protein was upregulated during mitochondrial stress (Fig. 5C, D). 23

30

Moreover, COQ-1::GFP upregulation was dependent on atfs-1 activity (Fig. 5E) and the 1

GFP signal of this construct was specific to the coq-1 gene (Fig. S17A-D). These results 2

show that in addition to regulating HMGS-1, an enzyme in the main branch of the 3

pathway, the UPRmt, via afts-1 activation, can upregulate enzymes in specific sub-4

branches. We propose that the coordinated upregulation of HMGS-1 along with 5

enzymes of specific sub-branches is a genetic program that has evolved to facilitate a 6

protection from mitochondrial dysfunction. 7

8

Discussion 9

We have found that mitochondrial dysfunction activates the UPRmt signaling cascade to 10

remodel cellular metabolism through the coordinated control of critical regulatory points 11

in the mevalonate pathway. Three independent approaches—i.e., whole transcriptome 12

expression profiling (NARGUND et al. 2012), RNAi-based functional assays, and targeted 13

qPCR analyses— demonstrated an upregulation in mevalonate pathway enzymes. 14

These included the main branch enzyme HMGS-1, enzymes responsible for the 15

synthesis of the ETC electron carriers ubiquinone and hemeA, and an enzyme that 16

facilities geranylgeranylation of proteins. The upregulation of the HMGS-1 enzyme relies 17

on the activity of two transcription factors, ATFS-1 and DVE-1, previously shown to play 18

a critical role in the mitochondrial stress response. We propose that this genetic circuit 19

has evolved to upregulate the synthesis of electron carriers that could alleviate 20

mitochondrial dysfunction, while in parallel enhancing geranylgeranylation, which is 21

required for the full activation of the UPRmt response. 22

31

One of the established roles of geranylgeranylation is to anchor one type of G-1

proteins, the small GTPase, to the membranes of different cellular compartments 2

(VOGLER et al. 2008). Several lines of evidence suggest that small GTPases play a role 3

in the regulation of various stress responses (BAR et al. 2016). Therefore, we 4

speculated that the inhibition of UPRmt by a block in geranylgeranylation (Fig. 4D-F) 5

stems from the inactivation of a small GTPase that is required for the UPRmt. Our 6

targeted screen, however, did not result in the identification of such a small GTPase. 7

One possible interpretation of these results is that our screen either lacked the small 8

GTPase essential for UPRmt execution, or, due to the partial effect of RNAi knockdown, 9

missed this small GTPase. Alternatively, it is plausible that a functional overlap between 10

two or more small GTPase proteins masks a requirement of a single small GTPase for 11

the activation of UPRmt. This redundancy, however, does not hold in treatment with 12

statins that impair the activity of most, if not all, the small GTPases that need to undergo 13

geranylgeranylation for their activity. Another possibility is that geranylgeranylation of a 14

protein that is not a small GTPase is necessary for UPRmt execution. For example, the 15

lack of geranylgeranylation of the nuclear lamina protein lamin A, that normally is 16

subjected to this type of modification (DAVIES et al. 2009), may block the translocation of 17

proteins, including ATFS-1, into the nucleus. 18

The mevalonate pathway accounts for the synthesis of several biomolecules that 19

play a role in diverse cellular processes, including the regulation of membrane fluidity, 20

respiration, and post-translational modification of proteins. So far, most studies in 21

mammals have been focused on the regulation of HMGCR1 (BROWN AND GOLDSTEIN 22

1980; BURG AND ESPENSHADE 2011) or the control of enzymes in the cholesterol 23

32

synthesis sub-branch (GILL et al. 2011; FORESTI et al. 2013; PRABHU et al. 2016). 1

Because HMGCR1 is the proposed rate-determining enzyme of the pathway in 2

mammals, it might be regulated in other cases that require remolding of mevalonate 3

pathway metabolism for example in mitochondrial dysfunction. The lack of regulation of 4

C. elegans HMGR-1 in mitochondrial stress may represent a nematode-specific 5

property of HMGR-1 regulatory network. Another possibility, however, is that we have 6

discovered an evolutionally conserved mechanism by which the UPRmt controls the 7

HMGS-1/HMGCS1 protein rather than controlling the HMGR-1/HMGCR1 enzyme. The 8

molecular details of the UPRmt in mammals are not well characterized (HAYNES et al. 9

2013). Nevertheless, a recent study of combined transcriptomic and proteomic analyses 10

of human cells exposed to EtBr has identified HMGCS1 but not HMGCR1 among the 11

list of a hundred proteins whose levels are mostly affected by the UPRmt (BAO et al. 12

2016). In contrast to our findings in C. elegans, HMGCS1 was not upregulated but 13

downregulated upon UPRmt activation. While HMGCS1 and HMGS-1 seem to be 14

regulated in opposite directions, these two observations suggest an evolutionary 15

conserved mechanism in which the UPRmt governs the levels of the first dedicated 16

enzyme of the pathway, HMGS-1/HMGCS1. We thus propose that the regulation of 17

HMGCS1/HMGS-1 levels plays a central role in the regulation of mevalonate pathway 18

flux in normal physiology, in stress conditions, and probably in diseases of mitochondrial 19

dysfunction. 20

In mammals, SREBP-1 upregulates HMGCS1 under conditions of low cholesterol 21

(HORTON et al. 2003), whereas p53 upregulates HMGCS1 during tumorigenesis (FREED-22

PASTOR et al. 2012). We did not find a role for the respective C. elegans orthologs sbp-1 23

33

and cep-1 in the upregulation of HMGS-1 upon UPRmt (Fig. 2G). Our results suggest a 1

context-dependent regulation of HMGCS1/HMGS-1 proteins by transcription factors 2

dedicated to different physiological conditions. Thus, we predict that in humans the 3

regulation of HMGCS1 in mitochondrial stress does not rely on the activity of SREBPs 4

or p53. Instead, we propose that HMGCS1 is regulated by transcription factors 5

dedicated to the UPRmt response such as ATF4 (BAO et al. 2016) or ATF5 (FIORESE et 6

al. 2016). 7

In mammals and in C. elegans, whole proteome studies have identified specific 8

residues of HMGCS1/HMGS-1 proteins as undergoing phosphorylation (VAN HOOF et al. 9

2009), acetylation (CHOUDHARY et al. 2009), ubiquitination (KIM et al. 2011; WAGNER et 10

al. 2011; WAGNER et al. 2012), or SUMOylation (SAPIR et al. 2014). These findings 11

highlight the possibility that HMGCS1/HMGS-1 undergoes a complex post-translational 12

regulation. In support of these findings, we have identified SIRT2.2 and AMPK-1 in a 13

proteomic screen for HMGS-1 interactors in C. elegans (SAPIR et al. 2014). The 14

biological significance of these post-translational modifications and the relative 15

functional weight of transcriptional versus post-translational regulation of 16

HMGCS1/HMGS-1 proteins will be important topics for future investigations. 17

Aside from upregulating HMGS-1 in the main trunk of the pathway, we found that 18

C. elegans’ UPRmt also upregulates enzymes that play a role in geranylgeranylation and 19

the synthesis of electron carriers. The identification of M57.2, the ortholog of human 20

RABGGTA (Rab geranylgeranyltransferase alpha subunit), as a gene upregulated by 21

spg-7 RNAi (Fig. 5A) suggests that the cellular protection of UPRmt involves an increase 22

in geranylgeranylation. Because UPRmt activation requires a functional 23

34

geranylgeranylation sub-branch (Fig. 4F), M57.2 upregulation may facilitate a stronger 1

UPRmt response in cases of mitochondrial dysfunction. A functional mevalonate 2

pathway was shown to be requisite for the activity of miRNAs in C. elegans (SHI AND 3

RUVKUN 2012), raising the possibility that the block of UPRmt stems from the 4

dysregulation of miRNAs. Nevertheless, this requirement of mevalonate pathway 5

metabolism for the activation of miRNAs was mapped to the dolichol synthesis sub-6

branch (SHI AND RUVKUN 2012), which, unlike the geranylgeranyl sub-branch (Fig. 4F), 7

does not alter the pattern of UPRmt activation (Fig. S13). Importantly, knockdown of the 8

M57.2 gene results in the activation of the UPRer (MORCK et al. 2009). Thus, by 9

controlling the levels of M57.2 and consequentially the level geranylgeranylation, the 10

UPRmt response can positively regulate its own activation and the activation of the 11

UPRer response. 12

The upregulation of enzymes that are responsible for the synthesis of ubiquinone 13

and hemeA electron carriers may be part of a compensatory mechanism that attempts 14

to restore mitochondrial hemostasis. In yeast, a growing body of evidence suggests that 15

mitochondrial malfunction results in the activation of transcription factors that increase 16

the level of ubiquinone-synthesizing enzymes (review in (GONZALEZ-MARISCAL et al. 17

2014)). In humans, several endogenous or environmental conditions lead to ubiquinone 18

deficiency or overproduction. However, beyond the activity of PPARthe molecular 19

mechanisms that control these effects remain largely unknown (BENTINGER et al. 2010). 20

Because CoQ10 is the end product of the sub-branch that synthesizes ubiquinone in 21

humans, CoQ10 supplementation is a suggested treatment to circumvent the adverse 22

side effects of statins. Our study suggests that the upregulation of enzymes from the 23

35

sub-branches that produce ubiquinones, hemeA, and geranylgeranyl moieties 1

represents a compensatory mechanism that is activated when the levels of these 2

metabolites are reduced. Based on these data, we propose that to overcome the 3

adverse effects of statins, a clinical approach should focus on the activation of 4

geranylgeranylation and the upregulation of hemeA synthesis, along with 5

supplementation with CoQ10. Supporting this hypothesis is a report that suggests that 6

supplementation with either mevalonate or geranylgeranyl pyrophosphate, but not with 7

other metabolites of the pathway, can mitigate the anti-tumorigenic effects of statins in 8

culture (JIANG et al. 2014). This report also suggests that geranylgeranyl-transferase II 9

inhibition slows cancer growth, implying that impaired geranylgeranylation is a 10

mechanism underlying the anti-tumorigenic activity of statins. Our results show, 11

however, that blocking of geranylgeranylation inhibits the UPRmt and potentially blocks 12

the compensatory mechanism of ATFS-1 activation and mevalonate pathway 13

upregulation. 14

15

Author contributions 16

A.S. designed the research; O.O., S.L., I.L.G., and A.S. conducted the experiments and 17

analyzed the data; A.S. wrote the paper. 18

19

Acknowledgments 20

Some strains were provided by the CGC, which is funded by the NIH Office of Research 21

Infrastructure Programs (P40 OD010440). We are grateful to Drs. Cole Haynes (UMass, 22

Amherst, MA), David. L. Baillie (SFU, Burnaby, BC, Canada), Matt Kaeberlein (UWash, 23

36

Seattle, WA), Paul. W. Sternberg (Caltech, Pasadena, CA,), Shane L. Rea (UT, 1

Houston, TX), and Sivan Henis-Korenblit (Bar-Ilan U, Ramat Gan, Israel) for C. elegans 2

strains. We thank Drs. Yoram Gercheman, Elah Pick, and Mr. Benjamin Trabelsi and 3

Mr. Shamsuzzama (all from the University of Haifa, Haifa, Israel) for reagents and 4

assistance with the analysis of qPCR results. We thank Drs. Limor Broday (Tel-Aviv 5

University, Tel Aviv, Israel), Ayelet Lamm, and Benjamin Podbilewicz (The Technion, 6

Haifa, Israel) for providing reagents and lab space. We are grateful to Vinci Au, Pegah 7

Abyaneh, Mark Edgley, and Dr. Donald G. Moerman (The University of British 8

Columbia, Vancouver, BC, Canada) for generating, by the CRISPR-Cas9 method, the 9

hmgs-1 deletion allele. The generation of this deletion allele was supported by the CIHR 10

(Canadian Institute for Health Research) grant to Dr. Donald G. Moerman. This work 11

was supported by the Israel Science Foundation (ISF) grants 41764 and 41765 to A.S. 12

13

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9

Figure legends 10

Figure 1. Mitochondrial Stress Upregulates HMGS-1 11

A) HMGS-1::GFP protein in worms fed with an empty vector control (EV). B) spg-7 12

dsRNA (spg-7i) leads to HMGS-1::GFP upregulation. C) HMGS-1::GFP protein in 13

worms of an isp-1(+) background. D) The isp-1(qm150) mutation leads to HMGS-14

1::GFP upregulation. E) HMGR-1::GFP in worms fed with an empty vector control. F) 15

spg-7 dsRNA does not evidently affect the level or distribution of HMGR-1::GFP. G) 16

Expression of the hmgs-1 promoter fused to gfp in worms grown on an empty vector 17

control. H) spg-7 dsRNA activates the hmgs-1 promoter. I) qPCR analyses of the 18

relative levels of endogenous hmgs-1 and hmgr-1 transcripts in worms fed with spg-7 19

dsRNA and empty vector control. 20

Error bars represent standard errors. “**”p≤0.01; NS indicates no significant difference 21

using a two tailed, unpaired Student’s T-test. J) Western blot analysis showing HMSG-22

1::GFP upregulation upon mitochondrial stress induced by the atp-3 RNAi. Actin was 23

used as a loading control. The size of the primary band (~85 kDa) is in agreement with 24

the calculated size of HMSG-1::GFP (83.4kDa), based upon the amino acid sequence 25

of the fusion protein. In all subsequent images, groups of ten worms were oriented with 26

their anterior side to the left. Unless otherwise stated, in all subsequent experiments, 27

41

worms were analyzed at the same chronological age (either day one or two of 1

adulthood). In each experiment, control and experimental worms were synchronized 2

and analyzed at the exact same chronological age. isp-1 mutants worms were analyzed 3

six days after the L1 stage. Consistent with previous reports (NARGUND et al. 2012; 4

PELLEGRINO et al. 2014), differences in worm size stem from the induction of UPRmt that 5

results in a smaller body size of the adult worms. Scale bars represent 250m (A-H) 6

and 200m (inserts, A-H). 7

8

Figure 2. The UPRmt, by Activating ATFS-1 and DVE-1, Upregulates HMGS-1 9

A) A scheme demonstrating the molecular details of the mitochondrial stress response 10

in C. elegans. B) HMGS-1::GFP in isp-1(qm150) mutant worms. We used this 11

background for the targeted screen. C) hmgs-1 RNAi abolishes the GFP signal, 12

demonstrating the specificity of the GFP construct and the hmgs-1 RNAi. D-F) 13

Knockdowns of atfs-1 (D), dve-1 (E), and rheb-1 (F) decrease the level of HMGS-1 14

upregulation. G) The complete score of the targeted RNAi screen. In each condition 15

tested, n (number of worms)= 100-120 worms. For each dsRNA tested, the results are 16

presented as the percent of the total number of worms monitored. The screen was 17

performed by the analysis of live intact worms under a fluorescence dissecting 18

microscope that enabled the qualitative assignment of worms into the three different 19

categories. Scale bars represent 250m and 500m (inserts). 20

21

Figure 3. Deficiency of HemeA and Ubiquinone activates the UPRmt 22

42

A) The expression pattern of the UPRmt reporter hsp-60pr::gfp in worms fed an empty 1

vector control. B) Knockdown of the cox-15 enzyme, which acts in the synthesis of 2

hemeA, upregulates hsp-60pr::gfp expression. C) hsp-60pr::gfp in the background of an 3

atfs-1(tm4525) loss-of-function mutation while fed an empty vector control. D) In the 4

background of atfs-1(tm4525), cox-15 RNAi did not upregulate the hsp-60pr::gfp 5

reporter. E) The expression of the hsp-6pr::gfp reporter in wild-type worms. F) coq-6

1(ok749) loss activates of the UPRmt. (G) Activation of the UPRmt, upon coq-1 loss, is 7

dependent upon atfs-1, and dve-1 (H). I) A thrashing assay revealed that activation of 8

the UPRmt, via atfs-1, does not protect the worms from the effect of coq-1 loss. 9

n(number of worms)= 20 for each condition. Bars represent standard errors. Statistical 10

significance was tested by the one-way ANOVA test using the SPSS software, “***” 11

represents p<0.005, “N.S.” indicates no significant difference. The details of the 12

statistical analyses can be found in the material and methods section. Differences in 13

worm size stem from the effect of the mutant or RNAi backgrounds. The scale bars 14

represent 500m (A-D); 250m (E-H). 15

16

Figure 4. Activation of the UPRmt Requires Geranylgeranylation 17

A-F) The results of a targeted screen for the requirement of different branches of the 18

mevalonate pathway for UPRmt signaling. A) The isp-1(qm150), hsp-6pr::gfp , worm 19

strain that was used in the screen. B) Knockdowns of atfs-1 and dve-1 (C) were used as 20

positive controls. D) Knock-down of hmgs-1 or hmgr-1 (E) inhibit the UPRmt. F) Knock-21

down of the geranylgeranyltransferase ggtb-1 gene suppresses the UPRmt. G-J) A 22

constitutively activated ATFS-1 protein in the background of the UPRmt reporter hsp-23

43

60pr::gfp. G) An empty vector control. H) atfs-1 RNAi reduces the level of UPRmt 1

activation in most tissues. The expression in the pharynx may stem from the resistance 2

of this tissue to RNAi knockdown (KUMSTA AND HANSEN 2012). I) Knock-downs of hmgs-3

1 or ggtb-1 (J), although resulting in potent visible phenotypes, do not suppress the 4

upregulation of hsp-60pr::gfp by the activated ATFS-1 protein. Differences in worm size 5

stem from the effect of the different RNAis. The atfs-1 and the isp-1 mutant worms were 6

analyzed four and six days after the L1 stage respectively. The scale bars represent 7

500m. 8

9

Figure 5. The Mitochondrial Stress Response Upregulates Multiple Branches of 10

the Mevalonate Pathway 11

A) qPCR analyses of the relative transcription levels of enzymes from different branches 12

of the mevalonate pathway. The pathway metabolites are labeled in black. Enzymes 13

labeled in green were analyzed by qPCR; enzymes in gray were not analyzed. Gray-14

blue arrows represent the flow of metabolites from the main branch to the end products. 15

N2 worms grown on bacteria transformed with the empty vector and spg-7 dsRNA 16

plasmids were used for experiments. Dark-orange arrows mark enzymes that were 17

suggested to be upregulated upon mitochondrial stress in a whole transcriptomic 18

analysis (NARGUND et al. 2012). Error bars represent standard errors of the mean. 19

“*”p≤0.05; “**”p≤0.01; “***”p≤0.001 using two tailed, unpaired Student’s T-tests. B-E) 20

The levels and distribution of a COQ-1::GFP reporter. B) The distribution of the COQ-21

1::GFP protein in worms fed with an empty vector control. The COQ-1::GFP protein is 22

enriched in the worm’s intestine. C) RNAi for spg-7 upregulates COQ-1::GFP levels 23

44

primarily in the intestine. D) Dilution of the spg-7 dsRNA by half did not evidently affect 1

its ability to upregulate the COQ-1::GFP protein. E) atfs-1 RNAi blocks the upregulation 2

of COQ-1::GFP in the spg-7 RNAi background. Scale bars represent 200m. 3