Research Article

CC2D1A causes ciliopathy, intellectual disability,
heterotaxy, renal dysplasia, and abnormal CSF flow
Angelina Haesoo Kim1,* , Irmak Sakin2,3,* , Stephen Viviano1 , Gulten Tuncel4, Stephanie Marie Aguilera5,
Gizem Goles5, Lauren Jeffries6, Weizhen Ji6, Saquib A Lakhani1,6, Canan Ceylan Kose7 , Fatma Silan7,
Sukru Sadik Oner8,9 , Oktay I Kaplan10 , MarmaRare Group11,†, Mahmut Cerkez Ergoren12 , Ketu Mishra-Gorur13,
Murat Gunel13,14,15,16, Sebnem Ozemri Sag17, Sehime G Temel17,18,* , Engin Deniz1,6,*

Intellectual and developmental disabilities result from ab-
normal nervous system development. Over a 1,000 genes have
been associated with intellectual and developmental dis-
abilities, driving continued efforts toward dissecting variant
functionality to enhance our understanding of the disease
mechanism. This report identified two novel variants in
CC2D1A in a cohort of four patients from two unrelated
families. We used multiple model systems for functional
analysis, including Xenopus, Drosophila, and patient-derived
fibroblasts. Our experiments revealed that cc2d1a is
expressed explicitly in a spectrum of ciliated tissues, in-
cluding the left–right organizer, epidermis, pronephric duct,
nephrostomes, and ventricular zone of the brain. In line
with this expression pattern, loss of cc2d1a led to cardiac
heterotaxy, cystic kidneys, and abnormal CSF circulation
via defective ciliogenesis. Interestingly, when we analyzed
brain development, mutant tadpoles showed abnormal
CSF circulation only in the midbrain region, suggesting ab-
normal local CSF flow. Furthermore, our analysis of the
patient-derived fibroblasts confirmed defective ciliogenesis,
further supporting our observations. In summary, we
revealed novel insight into the role of CC2D1A by
establishing its new critical role in ciliogenesis and CSF
circulation.

DOI 10.26508/lsa.202402708 | Received 11 March 2024 | Revised 29 July
2024 | Accepted 30 July 2024 | Published online 21 August 2024

Introduction

Intellectual and developmental disorder (IDD) is the result of
abnormal early development of the central nervous system that
clinically presents with intellectual and adaptive functioning def-
icits in conceptual, social, and practical domains (1). Approximately
2% of the world’s population is affected by IDDs, which remains the
most common reason for referral to genetic analysis (2, 3, 4).
Pathogenesis of the disease is complex; over a 1,000 genes have
been linked to non-syndromic IDD, suggesting a broad spectrum of
molecular defects presenting with intellectual disorders (3, 5). To
close this gap in our knowledge, ongoing work is focused on the
functional analysis of the variants and a better understanding of
the disease mechanism.

The coiled-coil and C2 domain–containing protein 1A (CC2D1A)
(OMIM#610055) has been implicated in a wide range of pathways,
including NF-κB enhancer binding protein signaling (6, 7), phos-
phodiesterase activity (8), and regulation of serotonin 1A, dopamine
D2 receptor (8, 9, 10, 11), bonemorphogenetic protein (12), and Notch
signaling (13). In the cytoplasm, it acts as a scaffold protein in the
PI3K/PDK1/AKT pathway and centrosome cleavage, mediating
centriole cohesion during mitosis (14). In vivo studies in mice
showed that CC2D1A is widely expressed in the brain (15), and the
cc2d1a-deficient mice died immediately after birth, indicating an
essential role during early development (16). Although the condi-
tional brain-specific knockout mouse models showed cognitive
deficits (17) and autistic-like phenotypes when CC2D1A was lost in

1Department of Pediatrics, Yale School of Medicine, New Haven, CT, USA 2Department of ENT, Cambridge University Hospitals NHS Foundation Trust, Cambridge, UK
3Acibadem University School of Medicine, Istanbul, Turkey 4DESAM Research Institute, Near East University, Nicosia, Cyprus 5Department of Neurosurgery, Yale
University School of Medicine, New Haven, CT, USA 6Pediatric Genomics Discovery Program, Department of Pediatrics, Yale University School of Medicine, New Haven, CT,
USA 7Canakkale 18 March University, Faculty of Medicine, Department of Medical Genetics, Canakkale, Turkey 8Department of Pharmacology, Goztepe Prof. Dr. Suleyman
Yalcin City Hospital, Istanbul, Turkey 9Istanbul Medeniyet University, Science and Advanced Technologies Research Center (BILTAM), Istanbul, Turkey 10Rare Disease
Laboratory, School of Life and Natural Sciences, Abdullah Gul University, Kayseri, Turkey 11MarmaraRare Group, Istanbul, Turkey 12Department of Medical Genetics,
Faculty of Medicine, Near East University, Nicosia, Cyprus 13Department of Neurosurgery, Yale School of Medicine, New Haven, CT, USA 14Department of Genetics, Yale
University School of Medicine, New Haven, CT, USA 15Yale Program in Brain Tumor Research, Yale University School of Medicine, New Haven, CT, USA 16Department of
Neuroscience, Yale University School of Medicine, New Haven, CT, USA 17Department of Medical Genetics, Faculty of Medicine, Uludag University, Bursa, Turkey
18Department of Histology and Embryology and Health Sciences Institute, Department of Translational Medicine, Faculty of Medicine, Bursa Uludag University, Bursa,
Turkey

Correspondence: engin.deniz@yale.edu; sehime@uludag.edu.tr
*Angelina Haesoo Kim, Irmak Sakin, Sehime G Temel, and Engin Deniz contributed equally to this work
†MarmaRare Group members are listed in the below Appendix.

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glutamatergic neurons (18), similar to homozygous mice, they also
died after birth, indicating potential additional roles of CC2D1A
during embryonic development. An additional mouse study in-
volving conditional postnatal removal of CC2D1A specifically in the
forebrain revealed morphologically abnormal cortical dendrite
organization and reduced density of dendritic spines, resulting in
mice with deficient neuronal plasticity, spatial learning, and
memory alongside features of reduced sociability, hyperactivity,
anxiety, and excessive grooming (19).

Various phenotypes have been associated with CC2D1A, also
known as Freud-1, Lgd2, and Aki-1, in the medical literature, but not
all studies include detailed phenotyping. Consistent with data from
murine modeling, the most well-characterized human patients
have neurological phenotypes. Homozygous pathogenic variants,
largely putative null alleles, have been associated with autosomal
recessive non-syndromic intellectual disability (OMIM# 608443) (15,

20, 21, 22) (Table 1). Several groups also have identified recessive
and dominant CC2D1A variants with various predicted molecular
consequences in different cohorts of patients with autism spectrum
disorder (ASD), with or without intellectual disability or seizures (21,
23, 24, 25, 26, 27, 28, 29).

Interestingly, Ma et al recently reported patients with congenital
heart disease presenting with damaging mutations in the CC2D1A
gene (31). The group identified seven damaging exonic missense
variants of CC2D1A in six patients with congenital heart disease
consistent with heterotaxy using whole-exome sequencing and
recapitulated heterotaxy in the zebrafish model system (31). Het-
erotaxy is caused by defective left–right (LR) patterning, where one
or more organs are misplaced along the left–right body axis where
cilia play a pivotal role (41, 42).

Cilia are membrane-bounded projections from the cell surface
and can be either motile, beating to generate extracellular fluid

Table 1. Summary of CC2D1A variants and clinical features of the affected individuals.

Nucleotide Protein Reported phenotype Reference

c.347delA p.(Lys116Argfs*82) Autism spectrum disorder, intellectual disability, and
seizures Manzini et al (23)

c.380C>T p.(Pro27Leu) Mental retardation, autosomal recessive Xiong et al (30)

c.575C>T p.(Pro192Leu) Heterotaxy Ma et al (31)

c.748 + 1G>T (splice impacted) Autism spectrum disorder, intellectual disability, and
seizures Manzini et al (23)

c.811delG p.(Ala271Profs*30) Intellectual disability, non-syndromic McSherry et al (32)

c.1016delC p.(Thr339Lysfs*95) Developmental disorder Abolhassani et al (33)

c.1345G>A p.(Val449Met) Neurodevelopmental delay Jauss et al (34)

c.1517A>G p.(Gln506Arg) Heterotaxy Ma et al (31)

c.1552G>T p.(Glu518*) Autism spectrum disorder Iossifov et al (24)

Lim et al (25)

Turner et al (26)

Fu et al (27)

Zhou et al (28)

c.1591G>A p.(Glu531Lys) Congenital heart defects Meerschaut et al (35)

c.1595C>T p.(Pro532Leu) Heterotaxy Ma et al (31)

c.1610C>T p.(Ser537Leu) Mental retardation, autosomal recessive Kamil et al (36)

c.1641 + 1G>A (splice impacted) Intellectual disability Rashvand et al (37)

c.1647G>T p.(Pro549=) Autism Zhou et al (28)

c.1739C>T p.(Thr580Ile) Smith–Magenis syndrome–like disorder, Joubert-like
disorder Loviglio et al (38)

Tuncel et al (22)

c.2342G>A p.(Gly781Glu) Heterotaxy Ma et al (31)

c.2342G>T p.(Gly781Val) Intellectual disability, heterotaxy Wang et al (39)

Ma et al (31)

c.2520-1G>T (splice impacted) Autism spectrum disorder Sener et al (29)

c.2657G>A p.(Arg886His) Smith–Magenis syndrome–like disorder Loviglio et al (38)

c.2693delG p.(Gly898Valfs*45) Intellectual disability, non-syndromic Reuter et al (40)

c.2728G>A p.(Glu910Lys) Mental retardation, autosomal recessive Xiong et al (30)

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flow (43), or immotile, acting as signaling centers (44, 45). Both types
of cilia are upstream of diverse biological processes in human
physiology, cell signaling, and embryonic development. Thus, the
diseases featuring heterotaxy are often associated with defective
ciliogenesis and are therefore commonly termed ciliopathies. The
role of CC2D1A in ciliary biology and how that relates to intellectual
disability remain to be determined.

This report shows that CC2D1A is essential for ciliogenesis and
plays a pivotal role in multiple developmental processes regu-
lated by cilia. We broaden the clinical spectrum linked to the
CC2D1A gene by presenting a new clinical presentation, cystic
renal disease, and show that CC2D1A is essential for nephro-
genesis. To date, only six variants in CC2D1A have been reported in
ClinVar to be pathogenic or likely pathogenic in patients pre-
senting with intellectual disability (ClinVar), and only around two
dozen variants have been clinically reported in the Human Gene
Mutation Database.

Here, we report two additional nonsense novel variants. A novel
homozygous (c.1186C>T [p.Arg396*]) variant in the CC2D1A gene was
identified in two siblings with intellectual disability, autistic fea-
tures, renal cyst, and obesity. Such a constellation of comorbidities
has been previously included in ciliopathies (46). A second family
with another novel homozygous (c.1264C>T [p.Glu422*]) variant was
identified in two siblings presenting with intellectual disability,
seizure disorder, and multiple renal cysts.

We used three distinct strategies to understand the role of
CC2D1A in development and disease. (1) Using the frog Xenopus
tropicalis model system, we revealed that cc2d1a is expressed
explicitly in a spectrum of ciliated tissues, including the gas-
trocoel roof plate (GRP, left-right organizer), epidermis, pro-
nephric duct, nephrostomes, otic vesicle, and ventricular zone of
the brain. We showed that CC2D1A is localized to the base of the
cilia in the GRP monociliated cells and the epidermal multi-
ciliated cells (MCCs). Tadpoles developed abnormal left–right
patterning and cystic kidneys when we depleted cc2d1a using the
CRISPR/Cas9 system. Interestingly, when we analyzed brain
development, mutant tadpoles showed abnormal cilium-driven
CSF circulation, specifically in the midbrain region. Using im-
munohistochemistry (IHC) and scanning electron microscopy
imaging, we showed that the midbrain ependymal motile cilia
decorating the brain ventricle were locally disrupted. We also
generated fibroblasts from our patients and demonstrated
defective ciliogenesis. (2) We tested the specific patient variant
by rescue experiments where WT human CC2D1A RNA injection
rescued the left–right patterning defect. In contrast, mutant RNA
with the patient variant failed, indicating that the patient variant
is indeed disease-causing. (3) Finally, to determine how CC2D1A
affects social behavior because the null and brain-specific
knockout mice died shortly after birth, we used the Drosoph-
ila model and well-established assays to study social behavior
(47). Indeed, CC2D1A mutant fruit flies recapitulated the anti-
social behavior our patients presented with.

In summary, our report expands on the role of CC2D1A in normal
development and its associated disease, which includes intel-
lectual and developmental disability, congenital heart disease,
and cystic kidney disease, establishing its essential role in
ciliogenesis.

Results

Clinical summaries and genetic analysis

We identified four patients from two unrelated families in different
parts of Turkey presenting with different homozygous variants in
the CC2D1A gene. Detailed clinical descriptions are presented
in Table 1. CC2D1A is highly conserved across the species; both
mutations were in proximity (Fig 1). To summarize, in Family 1, a
16-yr-old female patient presented with symptoms of intellectual
disability (ID), obesity, and ASD. Similarly, her 12-yr-old male sibling
presented with ASD and intellectual disability but in addition had a
dysplastic and dysfunctional left kidney harboring multiple renal
cysts. The siblings were born to healthy, consanguineous parents of
Turkish heritage by normal vaginal delivery with unremarkable
birth history. Both siblings carried a novel homozygous c.1186C>T
(p.Arg396*) nonsense variant inherited from each of the hetero-
zygous parents (Fig 1A and C). In Family 2, eight children were born
to healthy, consanguineous parents of Turkish heritage. Both
parents had heterozygous CC2D1A variants confirmed with Sanger
sequencing. Of the eight children, two siblings, with no significant
prenatal history, were referred to our clinic because of severe
intellectual disability and seizure disorder. Both siblings carried a
novel homozygous c.1264>T (p.Glu422*) nonsense variant inherited
from each of the heterozygous parents (Fig 1B and C). One sibling, a
17-yr-old male, also demonstrated dysmorphic facies and was
diagnosed with ASD. The other sibling, a 25-yr-old female, in ad-
dition to severe intellectual disability and seizure disorder, pre-
sented with renal cysts, multiple small cysts on the right kidney, and
a 15-mm parapelvic–cortical left renal cyst. She was also identified
to have a de novo missense variant c.11257C>T (p.Arg3753Trp)
(NM_001009944) in PKD1 (polycystin 1), a gene known to play a role
in kidney development and lead to autosomal dominant polycystic
kidney disease (48). This de novo mutation showed 0% frequency in
gnomAD and extremely low frequency in other population data-
bases. Segregation analysis was performed for Family 2 (parents
and male sibling), which confirmed the PKD1 variant to be de novo
and carried only by the affected female sibling. The results were
confirmed with tissue and blood Sanger sequencing.

Cc2d1a knockdown leads to dysplastic kidneys

Two of our patients from two different families presented with renal
cysts. The renal USG of the male patient (Family 1—12 yo) at 8 mo
of age demonstrated the cortical renal cyst (four cystic lesions
with dimensions of 18 × 12.2 mm, 14.9 × 14.8 mm, 23.5 × 14.6 mm, and
9 × 11 adjacent to the upper pole of the left kidney were repor-
ted—Table 2), and the follow-up DMSA study at 2 yr of age dem-
onstrated dysplastic, dysfunctional left kidney (Fig 1D). The renal
USG of the female patient (Family 2—25 yo) showed multiple bi-
lateral cystic lesions (Fig 1E), significantly more aggressive than our
male patient’s phenotype.

We investigated the impact of cc2d1a depletion on renal de-
velopment using the frog Xenopus model system. To begin our
studies, we used the whole-mount in situ hybridization to un-
derstand the expression pattern of the cc2d1a during renal

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development. Indeed, at stage 28 (post-fertilization day 3 at 26°C),
cc2d1a expression was strong in the pronephros and pronephric
duct (Fig 1F). Specifically, the three punctuate patterns of cc2d1a

expression correspond to the nephrostomes, which are ciliated
peritoneal funnels that connect the celomic cavity to the nephron
(49, 50), and the precursors to the kidneys before organogenesis

Figure 1. cc2d1a knockdown leads to dysplastic kidneys.
(A, B) Pedigrees and clinical presentation of four individuals from two unrelated families with homozygous mutations in CC2D1A, presenting with intellectual disability,
autism, obesity, renal cysts, and/or seizures. Symbols are as follows: filled, affected; empty, unaffected; circle, female; square, male; yellow circle, 16-yr-old female from
Family#1; red square, 12-yr-oldmale from Family#1; purple circle, 15-yr-old female from Family#2; green square, 17-yr-oldmale from Family#2; double bars, consanguinity in
the family; hash, deceased. Abbreviations: yo, year old. (C) Alignment of the CC2D1A amino acid sequence from the human, mouse, rat, and frog shows high conservation
across all species tested. The locations of the identifiedmutations are shown. (D) Renal ultrasonography of the 12-yr-old male patient from Family#1 at 8mo of age shows
cortical renal cysts (four cystic lesions). l, left; r, right; RPO, right posterior oblique; LPO, left posterior oblique. (E) Renal ultrasonography of the 25-yr-old female patient
from Family#2 shows bilateral cystic lesions. (F) Whole-mount in situ hybridization on WT X. tropicalis embryos at st.28 with antisense (left image) and sense (right
image) probes against cc2d1amRNA. a, anterior; p, posterior; d, dorsal; v, ventral; st., stage. (G) Zoomed-in image shows strong cc2d1a expression in the pronephric duct
and pronephros. Yellow arrows point to the nephrostomes, which are precursors to the kidneys. (H) Schematic of a one-of-two cell injection with cc2d1a sgRNA, Cas9, and
Dextran, Alexa Fluor 488 dye. The embryos were raised to st.46, and with fluorescencemicroscopy, we verified that only one side was injected (shown in green). The non-
fluorescent side serves as the internal control. r, right; l, left; a, anterior; p, posterior. (I) Representative 3D image of a st.46 Xenopus embryo, ventrally imaged using OCT.
The right side, depleted of cc2d1a, exhibited larger kidneys (as shown in the red circle). OCT, optical coherence tomography. (J) Quantification of kidney cross-sectional
area in the uninjected versus injected side in n = 28 embryos. Red and black dots indicate injected and uninjected side embryos, respectively. Paired t test; **** indicates
P < 0.0001.

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Table 2. Summary of CC2D1A high-confidence variants.

Pedigree Family 1 Family 2

Age (yr) 16 12 25 17

Gender Female Male Female Male

Birth 39 wk, vaginal delivery 39 wk, vaginal delivery

Geographic
origin Turkish Turkish Turkish Turkish

Parental
consanguinity + + + +

CC2D1A variant
c. 1186C > T c. 1186C > T c. 1264C > T c. 1264C > T

p. (Arg396*) p. (Arg396*) p. (Glutamine422*) p. (Glutamine422*)

PKD1 variant — — c.11257C>T (p.Arg3753Trp) —

Clinical
Characteristics

Intellectual
disability Severe Severe Severe Severe (IQ:30–39)

Renal cysts —

USG: a 14-mm-diameter cortical
cyst and a 10-mm-diameter
septate cortical cyst were
observed in the upper pole of the
left kidney.

USG: a few millimeter-sized
simple cortical cysts were
observed in the right kidney.
There are several thin-walled,
parapelvic/cortical-located
anechoic cysts in the left kidney,
the largest of which measures
15 mm in diameter in the middle
part, some of which have
calcified walls.

—

CT abdomen: 1. dense
calcification was observed in the
right adrenal gland location; 2.
cystic lesions were detected in
the upper pole of the left kidney.

DMSA renal scan: the right kidney
is in normal localization, normal
size, and smooth contours, and
shows homogeneous activity
within normal limits. The left
kidney is smaller in size and
smooth-shaped than the right,
and has a homogeneous, slightly
lower radiopharmaceutical
uptake compared with the right.
The contribution of the right
kidney to total kidney functions
was calculated as 63%, and the
contribution of the left kidney
was calculated as 37%.

Obesity + — — —

Autism
spectrum

Stereotypical movements,
echolalia+ + — +

Seizure
disorder — — Generalized tonic–clonic

seizures
Generalized tonic–clonic
seizures

Development
Expressive language difficulties,
occasionally unresponsive to
verbal stimuli at age 4.

Eye contact at 1.5 yr old.

Started walking at 4 yr old.
Started using single words at 4 yr
and forms non-coherent single-
and two-word sentences.

Started walking at 2 yr old. Non-
coherent words.

Other Obesity (> 97 percentile). Height
in 25–50 percentile.

Overweight, moderate persistent
asthma, cafe-au-lait single
submental lesion, hypermobility
of joints.

Length: < 0.02 percentile.

Dysmorphic facies: beaked nose,
prominent chin, thick eyebrows,
deep-set eyes. Nail hypoplasia.

Head circumference: < 5th
percentile.

Dysmorphic facies:
micrognathia, beaked nose, long
philtrum, thin narrow mouth

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(Fig 1G, yellow arrows). To analyze the impact of cc2d1a depletion on
renal development, we used a two-cell injection strategy and
depleted cc2d1a using the CRISPR/Cas9 system and quantified
post-editing using the ICE (Inference of CRISPR Edits) algorithm (51)
(Fig S1A and B). Cc2d1a CR#1+CAS9 +Dextran, Alexa Fluor 488 was
injected into one of two cells of embryos at the two-cell stage. With
this strategy, the uninjected side served as the internal control (Fig
1H). The embryos were then grown to st.45 (post-fertilization day 4
at 26°C) and visualized with optical coherence tomography (OCT)
imaging, where we acquired the 3D scan of the entire tadpole in vivo
(Fig 1I), as our group previously described (52). We measured the
largest cross sections of the kidney for both sides. In comparison,
the side of the embryo depleted of cc2d1a had overall larger,
dysplastic kidneys than the control side (Fig 1I and J). Together with
the observation of cc2d1a expression in the nephrostomes, and
depletion of cc2d1a leading to kidney dysplasia, these findings
closely recapitulated the kidney phenotype of our patients.

Cc2d1a mutant fruit flies recapitulated the antisocial behavior

To determine how CC2D1A affects social behavior because the null
and brain-specific knockout mice died shortly after birth, we used
the Drosophilamodel to study social behavior (47). We used a well-
established quantitative behavioral assay that tests Drosophila
social interaction, one of the three core ASD phenotypes as defined
by Diagnostic and Statistical Manual for Mental Disorders, fifth
edition (DSM-5; American Psychiatric Association, 2013). One of the
hallmarks of human ASD is the lack of proper interaction with other
individuals, which includes inappropriate responses to social cues,
causing them to either violate another person’s “personal space” or
overreact when another individual invades their personal space.
Hence, we used the assay for fly social spacing (analogous to
human social reciprocity), which exploits the natural tendency of
flies where, when housed as a group, flies settle into a “comfort-
able” social spacing that can be quantified using the social space
triangle (47).

Using the UAS-Gal4 system and actin-gal4 to drive the ubiquitous
expression of l(2)gd1 RNAi (l(2)gd1-IR), we assessed the functional
consequence of disrupting the lethal (2) giant discs (Lgd), the
Drosophila ortholog of CC2D1a, on the control of social spacing
behavior and the social space index (SSI) computed (47) (an SSI score
of ≤0 suggests little or no social interaction). The social space was
quantified for males, females, andmales plus females, and we found
a significant impact on the SSI in all three combinations as compared
to isogenic controls supporting the role of CC2D1A in ASD (Fig S2A–C).

Cc2d1a is required for proper left–right patterning

When we knocked down cc2d1a in Xenopus to examine its role
during development, one of the striking impacts was the disruption
of proper heart formation, often leading to the early demise of the
tadpoles. A critical step during heart development is the looping
process, when the tubular heart twists and loops around, forming
the chambers of the heart. Under normal conditions, the tubular
heart loops to the right, and this process relies on the proper LR
signaling of the body axis. In Xenopus, the cardiac sac is trans-
parent, allowing us to examine the cardiac looping and differentiate

a normally D-looped heart (dextra-looped—to the right) from ab-
normal phenotypes L-looped (levo-looped) or an A-looped
(ambiguous-looped) heart, where the outflow tract twists to the left
or has an indeterminate midline position (Fig 2A). When we knocked
down cc2d1a and raised the embryos to st.46 (post-fertilization day
4 at 26°C), we noticed that hearts were abnormally looped (Fig 2B).
Respectively, we used two non-overlapping CRISPRs targeting exon
1 (CRISPR#1) and exon 2 (CRISPR#2), which displayed 24% and 17%
looping defects (Figs 2B and S1A and B). Because proper heart
looping relies on proper LR patterning, we examined an upstream
marker of LR asymmetry, homeobox gene pitx2, that emanates from
the left lateral plate mesoderm in chick, mouse, and Xenopus (53,
54, 55, 56) (Fig 2C). When we knocked down cc2d1a, 26% (CR#1) and
22% (CR#2) of the embryos displayed the abnormal expression of
pitx2 at st.28 (post-fertilization day 3 at 26°C) as reversed, bilateral,
or absent, suggesting that the lateral plate mesoderm did not
correctly receive LR signaling (Fig 2D). We then examined the
singling upstream to pitx2 generated by the GRP.

The GRP is a ciliated structure that transiently forms at the dorsal
layer of the blastula, analogous to the Kupffer’s vesicle in zebrafish
and the node in mice and humans, and is responsible for estab-
lishing LR body axis patterning (57) (Fig 2E). We first asked whether
cc2d1a was expressed in the GRP, and using ISH, indeed, we
revealed that it was (Fig 2E–G). We then examined the expression of
the LR marker dand5 to analyze the impact of cc2d1a knockdown on
GRP patterning. During development, dand5 is a nodal antagonist
initially present symmetrically on the GRP at stages 14–16 (Fig 2H).
Then, on the surface of the GRP, motile cilia emerge and start to
beat at st.18–19, leading to a right-to-left fluid flow (gray arrows),
causing dand5 expression to be reduced on the left side of the
embryo (Fig 2I). This asymmetric inhibition of dand5 then activates a
signaling cascade downstream that leads the transcription factor,
pitx2, to be up-regulated on the left lateral plate mesoderm (Fig 2I),
setting the proper left–right axis (58, 59, 60, 61). Interestingly, we
observed the abnormal expression of dand5 at post-flow st.18, with
32% (CR#1) and 32% (CR#2) of embryos having bilateral, reversed, or
reduced/absent signal; however, we noticed a normal expression of
dand5 at pre-flow st.15 (Fig 2J). These findings suggested that cc2d1a
depletion results in a reduction of the dand5 inhibition on the left
side of the embryo, hinting at the possibility that cilium-driven flow
might have been compromised when cc2d1a is depleted. Before we
further investigated the potential impact on GRP cilia, we asked
whether the cardiac defect is specific to the cc2d1a knockdown via a
rescue experiment.

WT CC2D1A rescues abnormal LR patterning, and the patient
variant is detrimental to protein function

Our data so far suggested that cc2d1a regulates left–right pat-
terning; therefore, we examined cardiac looping in our rescue
experiments. We injected CR#1 and Cas9 protein at the one-cell
stage, followed by a co-injection with WT CC2D1A human RNA, and
raised these tadpoles to stage 46 (day 4) to score for cardiac
looping. The looping phenotype improved by 15% (Fig 2K). We, in
parallel, also tested the co-injection of the GFP-tagged human
CC2D1A RNA, which also rescued looping by 50%. However, intro-
ducing the patient variant RNA (c.1186C>T [p.Arg396*]) to cc2d1a-

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Figure 2. cc2d1a is required for proper left–right patterning.
(A, B) Representative images of cardiac outflow tract morphology and looping phenotypes in ventrally imaged st.45 Xenopus embryos. cc2d1amutants with abnormal
looping (L-loop and A-loop) are shown in red. Quantification of % cardiac looping defects in uninjected control (UIC) embryos, Cas9-only control, and cc2d1a CRISPR#1 and
CRISPR#2. CR#1- and CR#2-injected embryos displayed 24% and 17% looping defects, respectively. Data are shown as the mean ± SEM. Black dots represent individual
experiments, and the number of embryos is indicated above columns. P < 0.001 (***), P < 0.0001 (****). a, anterior; p, posterior; l, left; r, right; D-loop, dextra-looped; L-
loop, levo-looped; A-looped, ambiguous-looped; G0, generation 0; UIC, uninjected control; CR, CRISPR. (C) Whole-mount in situ hybridization of st.28 control embryos
showing the pitx2c signal (black arrow) on the left lateral plate mesoderm. (D) Representative images of st.28 cc2d1a mutant embryos with abnormal pitx2c expression
patterns (bilateral, reversed expression, or absent signal). Red arrows indicate the location of the present or missing pitx2c signal. CR#1- and CR#2-injected embryos
displayed 26% and 22% abnormal pitx2c expression, respectively. P < 0.01 (**). (E, F) Whole-mount in situ hybridization images of dissected GRPs from st.18 Xenopus

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depleted embryos failed to restore proper heart looping (Fig 2K).
Instead, the percentage of embryos co-injected with CR#1 and the
patient variant RNA observed were close to 25% abnormal looping,
suggesting that the patient variant is detrimental to function (Fig
2K). To further examine the role of cc2d1a in LR patterning, we
turned to the GRP cilia.

Cc2d1a is expressed at the GRP and localized to the base of the
monocilia, and its depletion resulted in abnormal cilia

Briefly, cilia are evolutionary-conserved, centriole-derived,
microtubule-based organelles protruding from the apical mem-
brane of the cells. There are three types of cilia: immotile monocilia
(sensory), motile monocilia, and motile multicilia. Given the results
so far, cc2d1a-depleted embryos exhibited loss of dand5 inhibition,
which relies on the presence of motile monocilium-driven fluid
flow. When we knocked down cc2d1a and analyzed the GRP cilia
with IHC, we observed abnormal monocilia (Fig 3A and B). Anti-
Arl13b antibody was used to label themonocilia, and phalloidin was
used to label the actin to mark cell borders. Indeed, GRP cilia
morphologically looked short and were depleted when cc2d1a was
knocked down. The average size of the GRP was unchanged, yet the
cilia per area were less (Fig 3C and D). When we injected N-GFP-
CC2D1A RNA to investigate the localization, we found that cc2d1a
was localized to the base of the GRP monocilia (Fig 3E and F). We
then proceeded to determine whether cc2d1a ciliary localization
and function are restricted to the GRP monocilia or are common to
other cilium types. We first turned to the motile MCCs on the
Xenopus epidermis.

Cc2d1a is localized to the base of the epidermal multicilia, and its
depletion leads to the loss of cilia; however, basal bodies were
preserved

The epidermis of the Xenopus is populated with MCCs analogous to
the human respiratory tract. We asked whether epidermal cilia are
also regulated by cc2d1a. First, similar to our findings in the GRP, the
N-GFP-CC2D1A localized to the base of the cilia on the MCCs. Next,
we labeled the rootlets to better understand the localization in
both GRP and epidermis. Rootlets are the cytoskeletal structure
that originates from the centrioles and anchors the basal bodies of
the cilia to the cell. We co-injected N-GFP-CC2D1A and CLAMP-RFP
(calponin homology and microtubule-associated protein to label
rootlets (62)) to visualize both proteins. CC2D1A was localized to the
tip of the ciliary rootlets, showing the exact localization in both GRP

monocilia and epidermal multiciliated cells (Figs 3G–I and S3A–F). To
determine whether the multicilia were abnormal like the GRP cilia,
we again used two-cell injections to deplete cc2d1a on the one
side. The embryos were grown to stages 28–30; then, we used OCT
to visualize cilium-driven flow along the epidermis as previously
described (63). Then, using IHC, we labeled the cilia with an anti-
acetylated tubulin antibody. We confirmed the loss of epidermal
cilia and cilium-driven fluid flow on the injected side with both
modalities (Video 1—Fig S4A–C). To further explain this loss of
ciliary phenotype, we investigated the centrioles at the base of the
cilia, referred to as basal bodies. Basal bodies are modified
centrioles that act as the microtubule organizer to form the cilia
and are localized at the tip of the rootlets, identical to the cc2d1a
localization. When we used IHC to co-stain the cilia with anti-
acetylated tubulin and the basal bodies with anti-gamma tubulin
in our cc2d1a-depleted tadpoles, we observed a loss of cilia but
did not appreciate a loss or disorganization of the basal bodies,
suggesting that despite defective ciliogenesis, centriole dupli-
cation, apicobasal migration, and proper membrane docking of
the basal bodies remained intact (Fig S4D).

Given that cc2d1a depletion led to two discrete types of ciliary
loss, GRPmonocilia and epidermalmulticilia, an intriguing question
is how these findings might be relevant to the intellectual disability
and autism spectrum disease that our patients and others in the
literature are presenting with. For this reason, we examined the cilia
in tadpole’s central nervous system.

Cc2d1a knockdown causes loss of cilium-driven CSF circulation in
the midbrain

In the Xenopus tadpole brain, cc2d1a expression showed a specific
location along the diencephalon and mesencephalon transition
zone, encapsulating the cerebral aqueduct (Fig 4D and E). In-
terestingly, when we analyzed the cilium-driven CSF circulation,
cc2d1a depletion led to the loss of CSF circulation in this specific
region.

Ependymal cilium-driven CSF circulation can be visualized in
Xenopus by OCT imaging. We have previously shown that the entire
Xenopus brain ventricular system can be visualized, and CSF flow
can be mapped using OCT imaging (64, 65). For this analysis, we
obtained in vivo optical midsagittal cross sections of st.46 Xenopus
brains to visualize brain morphology in WT and cc2d1amutants (Fig
4A). The Xenopus brain has four ventricles: lateral ventricle (tel-
encephalic), third ventricle (diencephalic), midbrain ventricle
(mesencephalic), and fourth ventricle (rhombencephalic). We have

embryos with sense (E) and antisense (F) probes against cc2d1a RNA. The black and white dotted area delineates the GRP, a ciliated structure that helps establish
left–right patterning. GRP, gastrocoel roof plate. (G) GRP cross section shows cc2d1a expression in the somites and notochord. GRP, gastrocoel roof plate; som, somites; no,
notochord; v, ventral; d, dorsal. (H) At st.15, an upstream LR patterning marker, dand5, is present symmetrically on both sides of the GRP. Motile cilia (green lines) emerge
and start to beat from right to left around st.18–19. The gray arrows indicate the fluid flow generated by the cilia. (I) Leftward fluid flow inhibits dand5 expression on the
left side of the GRP, leading to up-regulated pitx2c, which results in correct left–right asymmetry/organ situs. (J)Whole-mount in situ hybridization images of st.15 pre-flow
GRPs (top panel) show bilateral dand5 expression in cc2d1a mutant embryos, resembling the UICs. St.18 post-flow (bottom panel) cc2d1a mutant GRPs show bilateral
dand5 expression unlike the asymmetric expression shown in UICs. CR#1 and CR#2 GRPs both showed 32% abnormal post-flow dand5 expression. ns, not significant; P <
0.01 (**). (K) Quantification of % abnormal heart looping in UICs, CR#1 knockdown, human WT CC2D1A mRNA (10 pg), CC2D1A c.1186C>T p.Arg396* human patient variant
mRNA (10 pg), and human N-GFP WT CC2D1A mRNA (10 pg). cc2d1a mutant heart looping defects are rescued by human WT CC2D1A mRNA (15% rescue) and N-GFP WT
CC2D1AmRNA (50% rescue). The rescue fails with the patient variant mRNA (25% abnormal looping). Data are shown as themean ± SEM. ns, not significant; P < 0.05 (*), P <
0.01 (**), P < 0.001 (***). pg, picograms; WT, wild type; GFP, green fluorescent protein; hum, human.

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Figure 3. CC2D1A is expressed at the GRP and localized to the base of the monocilia, and its depletion results in abnormal cilia.
(A, B) Immunofluorescence images of control (A) and cc2d1a-depleted GRPs (B) stained with anti-Arl13b antibody (green) andmerged with phalloidin (purple). Scale bar
= 50 μm. (C, D) Quantification of the GRP size and (D) number of cilia per GRP area. Compared with the control, cc2d1amutant embryos showed no significant difference in
the GRP size, but did have fewer GRP cilia per area. ns, not significant; P < 0.0001 (****). (E) Immunofluorescence images of GRP (outlined in white) expressing human N-GFP
CC2D1A (green) stained with anti-Arl13b (red). (F) Zoomed-in images of GRP. Human N-GFP CC2D1A is expressed in the base of the monocilia. Scale bar = 25 µm.
(G) Immunofluorescence images of the epidermis of st.28–30 embryos expressing human N-GFP CC2D1A (green) stained with anti-AcTub (red) to show cilia. Human N-GFP-
CC2D1A localizes to the base of cilia in multiciliated cells (MCCs). The rightmost merged image is a close-up of a MCC. Scale bar = 10 μm. (H, I) St.28–30 embryos expressing
human N-GFP-CC2D1A (green) and CLAMP-RFP (red), which marks the rootlets. (H, I) Merged images show CC2D1A localizes to the tip of the ciliary rootlets in both GRP
monocilia ((H); scale bar = 5 μm) and epidermal MCCs ((I); scale bar = 15 μm).

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Figure 4. CSF flow is regionally compromised in cc2d1a mutant embryo brains with defective cilia.
(A, B)OCT-capturedmidsagittal section of st.46 control (A) and cc2d1amutant (B) tadpole brains. White lettering and dotted white lines indicate brain regions; the brain
ventricular system is labeled in yellow, and pink arrows indicate the location of the cerebral aqueduct. Ventricular CSF circulation was determined by particle tracking
map in the control st.46 tadpole brain and the cc2d1amutant tadpole brain. Green arrows indicate clockwise flow in the FF1 region and the FF4 region, whereas blue arrows
indicate anti-clockwise fluid flow in FF2, FF3, and FF5. Compromised CSF flow in the cerebral aqueduct is indicated by the red X. Scale bar = 50 μm. Temporal color coding
illustrates particle trajectory over time. The color scale bar shows the correspondence between the color and frame number in the color-coded image. a, anterior;
p, posterior; d, dorsal; v, ventral; Lat-V, lateral ventricle; III, third ventricle; M, midbrain ventricle; IV, fourth ventricle. (C)Quantification of CSF flow velocities in UIC, Cas9
control, and cc2d1amutant L-looped tadpoles,measured in μm/s, indicates slower FF2 and FF3 (cerebral aqueduct region) velocities in cc2d1amutant L-looped tadpoles. ns,
not significant; P < 0.05 (*), P < 0.0001 (****). CA, cerebral aqueduct; CSF, cerebrospinal fluid. (D, E)Whole-mount in situ hybridization on dissected WT X. tropicalis embryo
brains at st.45 with antisense (right image) and sense (left image) probes against cc2d1a mRNA (D). (E).Cc2d1a is expressed in the diencephalon and mesencephalon
transition zone, surrounding the cerebral aqueduct (E). a, anterior; p, posterior; d, dorsal; v, ventral; tel, telencephalon; di, diencephalon; mes, mesencephalon; rhom,
rhombencephalon; CA, cerebral aqueduct; st., stage. (F) Immunofluorescence images of dissected st.46 control stained with DAPI (top left in blue), anti-Arl13b (red, top
middle), and anti-γ-Tub (green, top right). The bottom shows merged images of Arl13b and γ-Tub. (G) SEM images of the ventral ependymal surfaces of st.46 control.
(H) Immunofluorescence images of dissected st.46 cc2d1amutant tadpole brains. In the cc2d1amutant brains, cilia in the cerebral aqueduct region are compromised,

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previously shown that each ventricle is decorated with ependymal
motile cilia and generates local flow fields (FFs)—FF1–5 (65). These
cilium-driven FFs have precise planar polarization and different
velocities based on location. Here, we marked them clockwise
versus anti-clockwise (Fig 4, Video 2). When we knocked down
cc2d1a, we didn’t observe a gross morphological change in the
brain. However, the FF2 and FF3 showed severely diminished or
near-absent CSF flow (Fig 4A–C and Video 2). The brain’s third
ventricle connects to the midbrain ventricle via the cerebral
aqueduct (purple arrow, Fig 4A). Similar to the mammalian brain,
aqueducts in Xenopus connect the ventricles and allow the
transport of molecules between the ventricles essential for
proper neurodevelopment. The CSF currents that regulate fluid
transport along the cerebral aqueduct are lost when cc2d1a is
depleted (Fig 4 and Video 2). The aqueduct where the CSF
currents are lost is located in the diencephalon–mesencephalon
transition zone where cc2d1a expression is enriched based on
our ISH data (Fig 4). Based on these findings, we next examined
the ciliary morphology of the ependymal surface. Flow fields 2
and 3 are localized to the ventral ependymal surface. We ana-
lyzed the ventral surface using IHC, where we marked cilia with
anti-Arl13b and basal bodies with anti-gamma tubulin, and also
used scanning electron microscopy for detailed analysis of the
ciliary morphology. Both analyses showed severely disrupted
cilia along the aqueduct (Fig 4G–I), explaining the loss of local
CSF circulation in this region.

We marked cilia with anti-Arl13b and basal bodies with gamma
tubulin, then analyzed the ventral surface using IHC. The analysis
showed severely disrupted cilia along the aqueduct (Fig 4F–H),
explaining the loss of local CSF circulation in this region.

Cc2d1a depletion led to defective cilia in Xenopus’s GRP, epi-
dermis, and ependyma. We finally asked whether patient-derived
fibroblasts demonstrated any ciliary defects.

Cultured fibroblast cells of patients demonstrate ciliary defects

Fibroblasts are mesenchymal cells of the connective tissue pro-
ducing the extracellular matrix and collagen, involved in wound
healing and scarring. When fibroblasts are cultured in low serum
medium for ~48 h, they form cilia (66, 67), and multiple works
demonstrated the in vitro use of fibroblasts to study ciliogenesis
(68, 69, 70). To obtain fibroblasts, skin punch biopsies were taken
from the patients and parents as described in the Materials and
Methods section. Western blot analysis of fibroblast cell line lysates
shows elimination of detectable CC2D1A expression in the two
patient fibroblasts compared with the control fibroblasts and the
father’s fibroblasts (Fig 5A). To analyze the cilia in fibroblasts, we
immunostained for acetylated tubulin to show cilia and phalloidin
to show the cytoskeleton. Under normal conditions, control fi-
broblasts showed a monocilium with an average length of 3.50 ±
1.15 μm (n = 590). In our index patients, either there were fewer
ciliated cells (Fig 5B) or the ciliary length was significantly dimin-
ished (Fig 5C–F).

Discussion

The CC2D1A gene has been identified in patients with a spectrum
of neurodevelopmental diseases, including non-syndromic auto-
somal recessive intellectual disability, ASD, and seizures, as well as
in patients with heterotaxy syndromes. Our work further expands
the clinical presentation and reports patients with CC2D1A
variants presenting with uni- and bilateral multicystic dysplastic
kidney disease. Although we must acknowledge the confounding
PKD1 variant in the 25-yo patient from Family 2 as a contributing
cause of her cystic kidney disease, we present evidence that
CC2D1A is also a contributing factor and could plausibly explain
such a severe kidney phenotype at such an early age, as most
patients with dominant PKD1 variants present as adults. Inter-
estingly, our analysis of cc2d1a expression during early devel-
opment revealed its association with nephrogenesis and the
ciliated structures. Cc2d1a is specifically expressed at the three
branches of the pronephric tubule (nephrostomes), known to be
densely ciliated, but this expression pattern is not limited to the
kidneys. Cc2d1a is highly expressed in the ciliated tissues
throughout early embryonic development. The GRP, epidermis,
nephrostomes, pronephric duct, optic vesicle, otic vesicle,
olfactory placodes, and ciliated ependymal surface of the
brain, specifically in the diencephalon and mesencephalon
regions of the brain, showed significant cc2d1a expression. We
also showed the cc2d1a localization at the base of mono- and
multicilia in discretely different cell types, suggesting a po-
tential global role in ciliogenesis. These findings align well
with the recent work from reference 31 where the authors
identified 26 probands with congenital heart disease, ex-
plicitly presenting with heterotaxy, which is well associated
with ciliopathy. The authors also showed that TALEN-induced
somatic cc2d1a knockdown in the zebrafish model recapitu-
lated the patient heterotaxy phenotype and showed defective
cilia in the central spinal canal (31). Of note, our brain findings
in Xenopus are different than the zebrafish findings in ref-
erence 31 report, where we see defective cilia in the midbrain;
however, the ependymal cilia in the hindbrain and spine re-
main unaffected.

Neurological defects are common in ciliopathies (71), and cilia
are known to play a critical role in cerebral cortex development (72).
Diverse neuropathologies in humans are associated with cilia,
including Joubert, Bardet–Biedl, Meckel–Gruber, and orofaciodigital
syndromes. It has been shown that cilia are involved in many
processes in neurodevelopment, including progenitor regulation
(73), interneuron migration (74), neural tube formation (75), and
cerebellar development (76). Recent advances also demonstrated
the role of cilia in embryonic CSF circulation (64, 65). This work
highlights a local, specific loss of CSF circulation in the midbrain
region when cc2d1a is lost, suggesting that cilia may also have
additional roles in regional brain development and function.
However, understanding the intricate relationship between local
cilium-based CSF circulation, brain development, and human

leading to slower CSF flow in FF2 and FF3 (light blue boxes). (I) SEM images of the ventral ependymal surfaces of st.46 cc2d1a mutant tadpole brains in the CA region
showing vastly reduced ciliary density. a, anterior; p, posterior; l, left; r, right; FF, flow field; CA, cerebral aqueduct; III, third ventricle; M, midbrain ventricle.

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neurocognitive disease remains incomplete. It will be important to
understand this relationship to define the pathophysiology better.

In summary, we have added evidence for CC2D1A as a cause of a
multisystem ciliopathy syndrome, expanding the spectrum of
CC2D1A-related disease from neurodevelopmental and cardiac
involvement to the currently included cystic renal disease, LR
patterning and CSF circulation. We also provide functional sup-
port for novel variants associated with this emerging disease and
introduce new understandings of ciliary biology as relates to
CC2D1A.

Materials and Methods

DNA sequencing and bioinformatics processing

Family 1
Written informed consent for participation in the study was
gained from both parents. Genomic DNA was extracted from

peripheral blood samples of patients and their parents using
QIAamp DNA Mini Kit (QIAGEN). The Clinical Exome Solution
(SOPHiA GENETICS) was used for exome enrichment. All proce-
dures were carried out according to the manufacturer’s protocols.
It is a capture-based target enrichment kit and covers 4,900 genes
with known inherited disease-causing mutations. Paired-end
sequencing was performed on an Illumina NextSeq 500 system
in Bursa Uludag University with a read length of 150 × 2. Base
calling and image analysis were conducted using Illumina’s Real-
Time Analysis software. The BCL (base calls) binary is converted
into FASTQ using the Illumina package bcl2fastq.

Bioinformatics analysis
All bioinformatics analysis was performed on the SOPHiA DDM
platform, which includes algorithms for alignment, calling SNPs and
small indels (Pepper), calling copy-number variations (Muskat), and
functional annotation (Moka). Raw reads were aligned to the hu-
man reference genome (GRCh37/hg19). Variant filtering and in-
terpretation were performed on SOPHiA DDM. Raw data were

Figure 5. Cultured fibroblast cells of patients demonstrate ciliary defects.
(A)Western blot of fibroblast cell line lysates from control, and Family#1 father, female patient, andmale patient. The affected patient cell lines did not show detectable
CC2D1A expression. (B, C) Quantification of fibroblast starvation–induced ciliary length. The patients displayed fewer ciliated cells or shorter ciliary lengths. ns, not
significant; P < 0.01 (**), P < 0.0001 (****). (D, E, F) Cultured and starved fibroblasts from Family#1 father, female patient, and male patient stained with DAPI (blue) to show
nuclei, phalloidin, Alexa Fluor 488 (green) to show actin cytoskeleton, and anti-acetylated tubulin (red) to show ciliary axonemes. Cilia on the patient fibroblasts were
less frequent and shorter than those on the control and father fibroblasts. Scale bar = 25 μm.
Source data are available for this figure.

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analyzed via the SOPHiA DDMdata analysis platform. Alignment and
variant discovery were performed by Pepper, a proprietary baseline
algorithm from SOPHiA GENETICS. Variant annotation was per-
formed with SOPHiA GENETICS’Moka software, and for each variant,
the effect of the variant on the protein sequence (missense, stop
gain, etc.), the frequency of occurrence in various populations
(1000G, ESP, ExAC, gnomAD), and prediction algorithms (SIFT, Poly-
Phen) were determined. Information such as the destructive effect of
the variant has been added. CNV detection was performed with
SOPHiA GENETICS’ Muskat software. Only variants located within
exonic regions and the 20-base pair border region between exons
and introns were included. Variants that passed the upstream
pipeline filters and with a call quality of ≥20 were included. Variants
with an allele frequency of >1% in GnomAD, 1000 Genomes, or ExAC
were excluded. Homozygosity mapping was carried out in families
with consanguineous marriages with HomSI (77).

Family#2
Written informed consent for participation in the study was gained
from both parents. Genomic DNA was extracted from peripheral
blood samples of patients and their parents using QIAampDNAMini
Kit (QIAGEN). Whole-exome sequencing analysis was performed
using the xGen Exome Research Panel v2 kit through the next-
generation sequencing method.

Bioinformatics analysis
The resulting Variant Call Format data were analyzed by creating
the following filter using QIAGEN Clinical Insight Interpret
8.1.20220121. Only variants located within exonic regions and the 20-
base pair border region between exons and introns were included.
Variants that passed the upstream pipeline filters and with a call
quality of ≥20 were included. Variants with an allele frequency
of >1% in GnomAD, 1000 Genomes, or ExAC were excluded.

Xenopus husbandry

X. tropicalis were housed and cared for in our aquatics facility
according to established protocols approved by the Yale Institu-
tional Animal Care and Use Committee. Animals were housed at our
aquatic facility under environmental control, including water
temperature, pH, and conductivity, as the stability of these variables
is essential. We followed the established protocol describing
conditions to optimally raise and maintain X. tropicalis from em-
bryos to adulthood (78).

Generation of the cc2d1a, pitx2, and dand5 probe and mRNA for
whole-mount in situ hybridization

The plasmids were linearized using the listed restriction enzyme
(SphI #R3182, Hind III #R3104, ClaI #R0197; NEB); then, the antisense
RNA probes were produced using HiScribe T7 High Yield RNA
Synthesis Kit (#E2040S; NEB) and DIG-dUTP (#03359247910; Roche).

Whole-mount in situ hybridization

In situ hybridization was performed on fixed embryos following
the standard protocol (80). However, the final fixation was done
with 4% PFA (#158127; Sigma-Aldrich) and 0.1% glutaraldehyde
(#5882; Sigma-Aldrich) in PBS (#P7059; Sigma-Aldrich) instead of
Bouin’s fixative. Expression patterns for pitx2c and dand5 were
assayed in stage 30 and stage 18 cc2d1a knockdown embryos,
respectively, and cc2d1a expression pattern was also assayed in
the WT stage 18 (GRP), stage 30 (epidermis, optic vesicle, otic
vesicle, nephrostomes, pronephric duct), and stage 45 (cranio-
facial structures). Removal of pigment by incubation in bleaching
solution (1% hydrogen peroxide [#H1009; Sigma-Aldrich] and 5%
formamide [#F7503; Sigma-Aldrich] in 0.5x SSC [#AB13156; Ameri-
can Bioanalytical]) was done after rehydration, before the 5-min
0.01 mg/ml proteinase K (#AB00925; American Bioanalytical)
treatment for stage 45 embryos. For all other stages, bleaching
was done after the final fixation after the BM Purple (#11442074001;
Sigma-Aldrich) color reaction.

CRISPRs, mRNA, and injections

CRISPR sgRNAs (small guide RNAs) for cc2d1a were designed from
the v9.0 model of the X. tropicalis genome using CRISPRscan (81):

CRISPR-1 (exon 1): 59-GGTCGGAAAGAAGTCCGTGGGGG-39.
CRISPR-2 (exon 15): 59-GCGCTGTTGTTTGGAGCGAAGGG-39.
sgRNAs were produced using EnGen sgRNA Synthesis Kit

(#E3322V; NEB). A pDONR221 plasmid containing a human CC2D1A
insert (reference sequence NM_017721) was obtained from
DNASU (#HsCD00829388). The insert was cloned into pCS DEST
(#22423; Addgene) and 223 pCS EGFP DEST (#13071; Addgene)
using Gateway cloning techniques (Invitrogen). The patient
variant (c.1186C>T) was produced using Q5 Site-Directed Muta-
genesis Kit (#E0554S; NEB) and also cloned into a pCS DEST
vector. mRNA was produced from these plasmids using mMES-
SAGE mMACHINE SP6 Transcription Kit (#AM1340; Invitrogen).
Xenopus embryos were produced by in vitro fertilization and
raised to appropriate developmental stages in 1/9x MR + 50 μg/
ml gentamicin (#G3632; Sigma-Aldrich) (82). Post-fertilization
embryos were injected at the one-cell or two-cell stage
according to standard protocols (82, 83, 84). 400 pg of CRISPR
sgRNA combined with 1.6 ng Cas9 protein (#CP03; PNA Bio) and a
fluorescent tracer, Dextran, Alexa Fluor 488 (#D22910; Invitrogen),
was injected at a volume of 2 nl into each embryo at the one-cell

Digoxigenin-labeled antisense mRNA probes against cc2d1a, pitx2 (58, 79), and dand5 (58, 79) were produced from the following plasmids.

Gene Sequence reference Expression vector Linearizing restriction enzyme

cc2d1a BC161089 pCS DEST Sph I

pitx2c TNeu083k20 pCS 107 Hind III

dand5 TEgg007d24 pCS 107 Cla I

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stage. For targeted loss-of-function experiments, 200 pg sgRNA
with 0.8 ng Cas9 protein was injected into one of two cells at the
two-cell stage; then, embryos were raised to the desired stages
(85, 86). For rescue experiments, 10 pg of human WT CC2D1A
mRNA, patient variant CC2D1A mRNA, or GFP-tagged CC2D1A
mRNA was injected separately into knockdown embryos at the
one-cell stage.

Genotyping/CRISPR analysis

To extract and purify the genomic DNA, stage 45 CRISPR–injected
embryos were dissociated in 50 μl of 50 mM NaOH (#7708-10;
Macron) at 95°C for 10 min (flicked every 3 min). Samples were
vortexed, then neutralized with 20 μl 1 M Tris (pH 7.4), and
centrifuged for 5 min. Supernatants were collected and stored
at −20°C. We designed primers around each CRISPR site for PCR
amplification using Primer3plus and then performed PCR using
Phusion High-Fidelity DNA Polymerase (#M0530S; NEB). The
primers used were CRISPR-1 (exon 1): 59-GAGCCCCCTGCATATAACCC-
39, 59-GGGCACTGCTATTCTAGTTGC-39, and CRISPR-2 (exon 15): 59-
CCTGGGACCTATTGCAAAGC-39, 59-CATCAGCACAGGAGCAAAGC-39. The
PCR products were run on an agarose gel; the fragments were gel-
extracted and purified using Monarch DNA Gel Extraction Kit
(#T1020S; NEB), then sent for Sanger sequencing (Quintarabio).
Sequencing results were then analyzed for cutting efficiency using
the Synthego ICE online tool. We verified that CRISPR/Cas9 edited
the proper cut site (Fig S1). Genotyping results to confirm
knockdown of the cc2d1a gene show a knockout score of up to 90%
for CRISPR#1 (targeting exon 1) and a knockout score of up to 81%
for CRISPR#2 (targeting exon 15). Overall knockdown over multiple
samples is represented in Fig S1.

Drosophila stocks and husbandry

Drosophila stocks were raised in standard food cornmeal/
molasses/agar bottles or vials at 25°C with a relative humidity of
20–40% in a 12-h dark–light cycle. l(2)gd1-IR, isogenic control, and
Actin-gal4 flies were obtained from the Bloomington Drosophila
Stock Center. All behavioral experiments were performed in a
genotype-balanced manner. To minimize the disruption of stan-
dard environmental conditions, flies were reared in bottles and
thus socially enriched, kept as mixed genders to allow mating, and
kept with standard food at all times before testing. Flies were
separated by gender the day before each experiment.

All experiments used flies naive to the test performed. Unless
otherwise noted in the text, the flies were collected from the bottles
when ~3–4 d old, sexed the day before the experiment, and placed
in vials (40/vial). Experiments were performed at the same time of
day from 11 am to 4 pm (between ZT4 and ZT9) to reduce variations
between trials. All behavioral assays were performed with a white
background using cardboard poster board and in a room at ~25°C
and ambient light.

Drosophila social space assay

The vertical triangle test chamber was constructed at the Yale
Machine Shop using the dimensions described by Simon, et al.

Briefly, vertical triangle test chambers were assembled using two
square glass plates (18 × 18 cm), separated by 0.47-cm spacers to
restrict flies within a 2D space (47, 87). Four spacers were used and
arranged into an isosceles triangle, with a 10-cm ruler placed on the
top-right surface for analysis scaling. Vials from the incubator were
allowed to acclimatize for 2 h before the experiment. The experi-
ment was conducted over 2 d under specific gender conditions:
male, female, and mixed (20_/20\), each with three independent
repeats. Flies were transferred from the vials to the chambers, and
after closing the entrance and securing the chambers with binder
clips, the chamber was tapped uniformly three times on a smooth
surface to standardize the starting position of the flies. Digital
images were captured every 30 min, three times per trial. Post-trial,
the flies were collected using the CO2 diffuser and returned to their
original vials for overnight recovery. This procedure was repeated
the next day at the same starting time for consistency. In total, 12
social behavioral assays were performed for each condition over
the 2 d.

Digital images were scaled using the in-frame ruler and con-
verted into Tagged Image File Format files using ImageJ, followed by
a batch conversion using Imaris software. Each fly was individually
identified using the spot selection tool, excluding any appearing
deceased. K-nearest neighbor distance was computed using Spots
(Imaris). Image-specific data were then exported to Excel and Prism
9 for further analysis, categorizing values into 0.5-cm bins. The
distribution of fly distances was visualized using binned
histograms.

The SSI was derived using the binned distance histograms. The
SSI was computed using the difference between the percentage of
flies in the first bin and the percentage of flies in the second bin. An
SSI below 0 indicated minimal to no social interaction among the
flies. Non-parametric tests, including the Kolmogorov–Smirnov and
Mann–Whitney tests, were applied to analyze the binned histo-
grams and SSI.

Xenopus cardiac looping

Post-fertilization stage 45 embryos were anesthetized in 2 g/liter
Syncaine (tricaine methanesulfonate; Syndel) in 1/9x MR and
ventrally scored for heart looping. The direction of heart looping
was determined by the position of the outflow tract (D-loop if
outflow tract curves to the right, L-loop if outflow tract loops to the
left, and A-loop if it does not loop).

IHC and imaging

GRP monocilia (st.18)
Control and cc2d1a knockdown embryos were raised to stage 18,
fixed with 4% PFA in PBS for 1 h at RT, and rinsed with PBS. GRPs
were dissected and incubated in a blocking buffer (3% BSA [#A9647;
Sigma-Aldrich] and 0.1% Triton X-100 [#AB02025 in PBS; American
Bioanalytical]) for 1 h at RT. Samples were then incubated with
mouse anti-Arl13b (NeuroMab clone N295B/66) diluted 1:100 in
blocking buffer overnight at 4°C. GRPs were washed with 0.1% Triton
X-100 in PBS three times for 10 min, incubated in blocking buffer for
30 min, and then incubated in donkey anti-mouse Alexa Fluor 594
(#A21203; Invitrogen) diluted 1:500 in blocking buffer at RT for 2 h.

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The samples were then washed with 0.1% Triton X-100 in PBS two
times for 10 min at RT. Actin filaments were stained with phalloidin,
Alexa Fluor 488 (#A12379; Invitrogen) diluted 1:100 in 0.1% Triton
X-100 in PBS at RT for 1 h; then, the samples were washed in PBS two
times for 10 min and mounted between coverslips with ProLong
Gold antifade mountant (#P36934; Thermo Fisher Scientific).

Epidermal multicilia (st.28–30)
The Xenopus epidermis is populated with multiciliated cells
allowing straightforward observation and functional analyses (88,
89, 90). Control and cc2d1a knockdown embryos were raised to
stages 28–30, then fixed and immunostained the same way as the
GRPs instead of using mouse anti-acetylated tubulin clone 6-11B-1
(#T6793; Sigma-Aldrich) and rabbit anti-γ-tubulin (#T3559; Sigma-
Aldrich) as the primary antibodies. Embryos weremounted between
coverslips in ProLong Gold using vacuum grease as a spacer.

Ependymal monocilia/multicilia (st.45)
Control and cc2d1a knockdown tadpoles were raised to stage 45.
Mutant tadpoles were scored for heart looping defects; then,
normal control and abnormally looped tadpoles were fixed with
4% PFA in PBS for 1 h and rinsed with PBS. To be able to better
observe brains, embryo heads were dissected, removing facial
cartilage, tail, gut, and lower jaw. The heads were then dehydrated
by washing twice with 100% methanol (#179337; Sigma-Aldrich)
and stored at −20°C overnight. The samples were then bleached in
10% hydrogen peroxide in 100% methanol at RT on direct light
until the pigment was sufficiently gone (about 3 h). After a rinse in
100% methanol, the samples were rehydrated stepwise (50%
methanol and 25% methanol, 10 min each) to TBS (155 mM NaCl
[#9888; Sigma-Aldrich] and 10 mM Tris [#AB02000; American
Bioanalytical], pH 7.5), then incubated in 0.1% Triton X-100 in TBS
overnight at 4°C. The next day, embryos were blocked in 10% FBS
(Sigma-Aldrich) and 0.3% Triton X-100 in BSDSGS (1% BSA, 5%
donkey serum [#017-000-121; Jackson ImmunoResearch], 5% goat
serum [#5425S; Cell Signaling Technology], 0.1% glycine
[#AB00730-01000; American Bioanalytical], 0.1% lysine [#AB145111;
Abcam] in PBS) for 4 h at RT, then incubated in primary antibodies
overnight at 4°C as described previously (89). The primary anti-
bodies used were rabbit anti-Arl13b (#17711-1-AP; Proteintech)
diluted 1:100 and mouse anti-γ-tubulin (#T6557; Sigma-Aldrich)
diluted 1:200 in 10% FBS, 0.1% Triton X-100 in BSDSGS. The embryos
were rinsed, then washed in TBST (TBS + 0.1% Triton X-100) at RT
for 1 h, washed in TBS three times for 1 h at RT, and washed in TBS
overnight at 4°C. The next day, they were incubated in secondary
antibodies, donkey anti-rabbit Alexa Fluor 594 and chicken anti-
mouse Alexa Fluor 488 (#A21200; Invitrogen) each diluted 1:500;
and Hoechst 33342 (#H3570; Invitrogen) diluted 1:5,000 in TBS for
2 h at RT. The samples were then washed in TBS three times for
10 min. Samples were then mounted between coverslips in Pro-
Long Gold using vacuum grease as a spacer.

Cultured fibroblasts
A 4-mm punch skin biopsy was taken under local anesthesia from
the patients (genotype confirmed) and an age-matched, unre-
lated, healthy, donor control. Written informed consent for par-
ticipation in the study was gained from the patients and control.

Sterile scalpel blades were used to cut the biopsies into smaller
pieces of 0.5 mm, which were then put into a six-well plate. The
tissue was incubated at 37°C in 5% CO2 in a humidified incubator
with a limited amount of growth media (DMEM, 10% FBS, 1 mM
sodium pyruvate, 4 mM L-glutamine, penicillin–streptomycin, and
2.5 μg/ml amphotericin B). Fresh medium (2 ml) was added the
next day. The fibroblasts were cultured for ~4 wk until enough
fibroblast outgrowth had occurred to allow for additional cell
passage (amphotericin B [J67049.AD; Thermo Fisher Scientific],
DMEM [#21068028; Gibco], L-glutamine [#25030081; Gibco], sodium
pyruvate [#11360070; Gibco], penicillin–streptomycin [#10378016;
Gibco], FBS [#10270-106; Gibco]). Passaging was done with 0.25%
trypsin–EDTA (#25200056; Gibco). Ciliary growth was induced by
incubating cultured cells for 48 h in growth medium without FBS.
Cells to be immunostained were cultured in eight-well culture
slides (#354108; Falcon/Corning), fixed in 4% PFA in PBS for 1 h, and
then washed three times with PBS. Cells were then incubated in
blocking buffer (3% BSA/PBS with 0.1% Triton X-100) for 1 h, then
incubated overnight at 4°C in a primary antibody (mouse anti-
acetylated tubulin; Sigma-Aldrich) diluted 1:1,000 in blocking
buffer. Cells were washed three times for 10 min with PBS, then
incubated in a secondary antibody (goat anti-mouse Texas Red,
#T6390; Invitrogen) diluted 1:500 in blocking buffer. Wells were
washed two times for 10 min with PBS, then incubated in phal-
loidin, Alexa Fluor 488 diluted 1:100 and Hoechst 33342 diluted 1:
5,000 in PBS for 30 min. Wells were then washed two times for
10 min with PBS. Then, wells were removed from the culture slide
and the cells were mounted with ProLong Gold.

All immunostained samples were imaged on a Zeiss LSM 880
confocal microscope. Fluorescent images were processed and
analyzed using Fiji/ImageJ (91). For GRP ciliary quantification, the
GRP area was outlined, and then, cilia were manually counted in
that region as described previously (79, 92). Fibroblast ciliary
lengths were measured using Zen (Blue Edition) version 3.6 soft-
ware (Zeiss).

Western blotting

To extract lysates from fibroblasts, cells were cultured in six-well
culture plates to ~80% confluence, then washed once with PBS,
and aspirated. SDS lysis buffer (2% SDS [#AB01922; American
Bioanalytical], 10% glycerol [#2136-03; Baker], and 62.5 mM Tris,
pH 6.8) was heated to 100°C; then, 150 μl was added to each well
of the six-well plate. The cells were lysed by swirling the slurry on
the bottom of the well with a pipet tip; then, the slurry was
transferred to microfuge tubes. The samples were incubated at
100°C for 10 min, cooled to RT, and then stored at −20°C until
needed. Samples were thawed on ice; then, protein concen-
trations were calculated using BCA Protein Assay Kit (#23225;
Thermo Fisher Scientific) according to the manufacturer’s in-
structions. 5 μg of each lysate with Laemmli sample buffer (#161-
0747; Bio-Rad) was run on a 4–12% Bolt Bis-Tris Plus gel
(#NW04120BOX; Invitrogen), then transferred to a PVDF mem-
brane (#1620219; Bio-Rad). The membrane was blocked for 1 h
with 5% non-fat dry milk (#AB10109; American Bioanalytical) in
TBST, then incubated overnight at 4°C in primary antibody
(mouse anti-CC2D1A, #H00054862-B01P; Thermo Fisher Scientific)

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diluted 1:250 in 5% non-fat dry milk in TBST. The membrane was
washed three times for 15 min in TBST, then incubated for 2 h in a
secondary antibody (donkey anti-mouse HRP, #715-035-150;
Jackson ImmunoResearch). The membrane was then washed
three times for 15 min in TBST. The SuperSignal West Pico Plus
chemiluminescent substrate (#34580; Thermo Fisher Scientific)
was used according to the manufacturer’s instructions to vi-
sualize stained protein bands. After exposure to the substrate,
the membrane was scanned using an Azure c300 Western blot
imager.

OCT imaging and CSF velocity quantification

OCT imaging was performed as we previously demonstrated (52, 63,
64, 65, 93). Stage 46 tadpoles were anesthetized in 2 g/liter Syncaine
in 1/9x MR, and cross-sectional (midsagittal) images of the brain
ventricles were obtained with OCT/ThorImage. 2D/3D images and 2D
Videos were used to quantify the brain areas and CSF flow velocities
using Fiji/ImageJ andMATLAB. The Gaussian process post-processing
was applied for particle velocimetry to quantify CSF flow velocity, as
we described previously (63). CSF flow was measured in μm/sec.
Figure images were built with averaged particle speed colorization
and were processed in Fiji, ImageJ (91). CSF circulation flow was
classified as normal, slow, or absent because of flow.

Scanning electron microscopy

The dissected tissue was fixed with 4% PFA overnight at 40°C,
followed by further fixation once the sample was pinned open
with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4
(#16537-20; Electron Microscopy Sciences), for 1 h. Samples were
rinsed in 0.1 M sodium cacodylate buffer (#J60344.AE; Thermo
Fisher Scientific) and post-fixed in 2% osmium tetroxide (#201030;
Sigma-Aldrich) in 0.1 M sodium cacodylate buffer, pH 7.4. These
samples were rinsed in buffer and dehydrated through an ethanol
series to 100%. The samples were dried using a Leica 300 critical
point dryer with liquid carbon dioxide as transitional fluid and
were glued to aluminum stubs using a carbon adhesive, and then
sputter-coated with 4 nm platinum 80/palladium 20 using
Cressington 208HR. The samples were viewed and digital images
acquired in Zeiss CrossBeam 550 between 1.5 and 2 kV at a working
distance of 8–12 mm.

Statistical analysis

All experiments had at least three replicates and were tested for
statistical significance using two-tailed t tests in GraphPad Prism 9.
Statistical significance was defined as P < 0.05 (*), P < 0.01 (**), P <
0.001 (***), and P < 0.0001 (****).

Appendix: MarmaRare Group

Yasemin Alanay, Yasemin Kendir-Demirkol, Ozlem Akgun Dogan,
Mahmut Cerkez, Ergoren, Ozden Hatirnaz Ng, Ugur Ozbek, Ozkan

Ozdemir, Sebnem Ozemri Sag, Ilayda Sahin, Sehime G Temel, Kanay
Yararbas.

Data Availability

De-identified data are available upon request from the authors.

Ethics declaration

All institutions involved in this research received approval from
their local IRB or Research Ethics Committee. Informed consent was
obtained from all individuals or their parents/legal guardians
through the IRB protocols at Yale University School of Medicine
(main IRB) or one participating institution. Individual data have
been de-identified; for the presentation of identifiable patient
images, express written consent has been obtained from the in-
dividuals or their parents/legal guardians. Animal research was
performed under an approved Institutional Animal Care and Use
Committee Protocol at the Yale University School of Medicine.

Supplementary Information

Supplementary Information is available at https://doi.org/10.26508/lsa.
202402708

Acknowledgements

The authors thank all the patients and their families for participating in our
research study. The authors thank the Yale Center for Genome Analysis for
DNA sequencing, and Xinran Liu and Morven Graham at the Yale Electron
Microscopy laboratory for assistance with micrographs. E Deniz was sup-
ported by NIH/NICHD R01NS127879.

Author Contributions

AH Kim: conceptualization, formal analysis, investigation, visuali-
zation, methodology, and writing—original draft, review, and
editing.
I Sakin: conceptualization, formal analysis, investigation, and wri-
ting—original draft, review, and editing.
S Viviano: conceptualization, resources, data curation, formal
analysis, validation, investigation, visualization, methodology, and
writing—original draft, review, and editing.
G Tuncel: data curation, formal analysis, validation, investigation,
and methodology.
SM Aguilera: investigation, visualization, and methodology.
G Goles: investigation, visualization, and methodology.
L Jeffries: data curation, formal analysis, and writing—review and
editing.
W Ji: data curation, formal analysis, validation, and writing—review
and editing.
SA Lakhani: data curation and formal analysis.
CC Kose: data curation and investigation.
F Silan: data curation, formal analysis, and investigation.

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https://doi.org/10.26508/lsa.202402708
https://doi.org/10.26508/lsa.202402708
https://doi.org/10.26508/lsa.202402708


SS Oner: investigation and writing—review and editing.
OI Kaplan: investigation and writing—review and editing.
MC Ergoren: data curation, formal analysis, investigation, meth-
odology, and writing—original draft, review, and editing.
K Mishra-Gorur: data curation, formal analysis, supervision, in-
vestigation, methodology, project administration, and wri-
ting—original draft, review, and editing.
M Gunel: data curation, formal analysis, supervision, investigation,
and project administration.
SO Sag: conceptualization, resources, data curation, formal anal-
ysis, supervision, validation, investigation, methodology, and wri-
ting—original draft, review, and editing.
SG Temel: conceptualization, resources, data curation, formal
analysis, supervision, funding acquisition, validation, investigation,
visualization, methodology, project administration, and wri-
ting—original draft, review, and editing.
E Deniz: conceptualization, resources, data curation, formal anal-
ysis, supervision, funding acquisition, validation, investigation, vi-
sualization, methodology, project administration, and
writing—original draft, review, and editing.

Conflict of Interest Statement

One author reports part ownership of startup companies unrelated to this
work: Qiyas Higher Health (SA Lakhani) and Victory Genomics (SA Lakhani).
All other authors declare no conflicts of interest.

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