Unleashed Actin Assembly in Capping Protein-Deficient B16-F1 Cells Enables Identification of Multiple Factors Contributing to Filopodium Formation

Abstract

Background:
Filopodia are dynamic, finger-like actin-filament bundles that overcome membrane tension by forces generated through actin polymerization at their tips to allow extension of these structures a few microns beyond the cell periphery. Actin assembly of these protrusions is regulated by accessory proteins including heterodimeric capping protein (CP) or Ena/VASP actin polymerases to either terminate or promote filament growth. Accordingly, the depletion of CP in B16-F1 melanoma cells was previously shown to cause an explosive formation of filopodia. In Ena/VASP-deficient cells, CP depletion appeared to result in ruffling instead of inducing filopodia, implying that Ena/VASP proteins are absolutely essential for filopodia formation. However, this hypothesis was not yet experimentally confirmed.

Methods:
Here, we used B16-F1 cells and CRISPR/Cas9 technology to eliminate CP either alone or in combination with Ena/VASP or other factors residing at filopodia tips, followed by quantifications of filopodia length and number.

Results:
Unexpectedly, we find massive formations of filopodia even in the absence of CP and Ena/VASP proteins. Notably, combined inactivation of Ena/VASP, unconventional myosin-X and the formin FMNL3 was required to markedly impair filopodia formation in CP-deficient cells.

Conclusions:
Taken together, our results reveal that, besides Ena/VASP proteins, numerous other factors contribute to filopodia formation.

Full text

Citation: Hein, J.I.; Scholz, J.; Körber,
S.; Kaufmann, T.; Faix, J. Unleashed
Actin Assembly in Capping
Protein-Deficient B16-F1 Cells
Enables Identification of Multiple
Factors Contributing to Filopodium
Formation. Cells 2023,12, 890.
https://doi.org/10.3390/
cells12060890
Academic Editor:
Annette Müller-Taubenberger
Received: 2 December 2022
Revised: 8 March 2023
Accepted: 10 March 2023
Published: 14 March 2023
Copyright: © 2023 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
cells
Article
Unleashed Actin Assembly in Capping Protein-Deficient B16-F1
Cells Enables Identification of Multiple Factors Contributing to
Filopodium Formation
Jens Ingo Hein, Jonas Scholz, Sarah Körber , Thomas Kaufmann and Jan Faix *
Institute for Biophysical Chemistry, Hannover Medical School, Carl-Neuberg-Strasse 1, 30625 Hannover, Germany
*Correspondence: faix.jan@mh-hannover.de; Tel.: +49-511-532-2928
Abstract:
Background: Filopodia are dynamic, finger-like actin-filament bundles that overcome
membrane tension by forces generated through actin polymerization at their tips to allow extension
of these structures a few microns beyond the cell periphery. Actin assembly of these protrusions is
regulated by accessory proteins including heterodimeric capping protein (CP) or Ena/VASP actin
polymerases to either terminate or promote filament growth. Accordingly, the depletion of CP in B16-
F1 melanoma cells was previously shown to cause an explosive formation of filopodia. In Ena/VASP-
deficient cells, CP depletion appeared to result in ruffling instead of inducing filopodia, implying
that Ena/VASP proteins are absolutely essential for filopodia formation. However, this hypothesis
was not yet experimentally confirmed. Methods: Here, we used B16-F1 cells and CRISPR/Cas9
technology to eliminate CP either alone or in combination with Ena/VASP or other factors residing
at filopodia tips, followed by quantifications of filopodia length and number. Results: Unexpectedly,
we find massive formations of filopodia even in the absence of CP and Ena/VASP proteins. Notably,
combined inactivation of Ena/VASP, unconventional myosin-X and the formin FMNL3 was required
to markedly impair filopodia formation in CP-deficient cells. Conclusions: Taken together, our results
reveal that, besides Ena/VASP proteins, numerous other factors contribute to filopodia formation.
Keywords: filopodia; capping protein; Ena/VASP proteins; myosin-X; mDia2; FMNL2; FMNL3
1. Introduction
The precisely coordinated spatiotemporal control of the assembly and disassembly of
actin filaments is a major determinant in a wide range of fundamental cellular processes
such as endocytosis, cytokinesis and cell migration [
1
–
4
]. Specific protein assemblies, com-
posed of numerous actin-binding proteins, act in these processes to nucleate, elongate or
cap new actin filaments, organize them into complex 3D arrays, and, subsequently, disas-
semble them to replenish the polymerization-competent pool of monomeric
G-actin [5,6]
.
Migration of cells on flat and rigid substrates is commonly initiated by the protrusion
of sheets of cytoplasm, referred to as lamellipodia, which are filled with dendritic actin
filament networks and compact filament bundles termed microspikes and filopodia, the
structure, dynamics and turnover of which have been extensively characterized [
7
–
10
].
The nucleation of branched actin networks in lamellipodia is driven by the actin-related
proteins 2 and 3 (Arp2/3) complex downstream of the WAVE-regulatory complex (WRC)
and Rac signaling [
11
–
14
]. The only protein families known so far to actively accelerate
the rate of actin filament elongation in the presence of CP (also known as CapZ in muscle)
by incorporating actin monomers at the growing barbed ends are formins and Ena/VASP
proteins, albeit their modes of action differ considerably [10].
Formins constitute a conserved group of large, multi-domain proteins that promote
the nucleation and elongation of unbranched (linear) actin filaments as found in stress fibers
(SF) and filopodia [
15
]. The crescent-shaped FH2 domain homo-dimerizes into a doughnut-
like structure [
16
], nucleates actin filaments and remains continuously associated with the
Cells 2023,12, 890. https://doi.org/10.3390/cells12060890 https://www.mdpi.com/journal/cells

Cells 2023,12, 890 2 of 25
progressively elongating barbed end, thereby effectively preventing termination of filament
growth by CP [
17
]. The FH2 domain is usually preceded by a formin homology 1 (FH1)
domain composed of consecutive stretches of poly-L-proline that serve as binding sites for
profilin–actin complexes, enabling recruitment and delivery of ATP-loaded actin monomers
to the FH2 domain for subsequent incorporation into growing filament barbed ends [
18
].
A subset of formins referred to as Diaphanous-related formins (Drfs) act as effectors of
Rho family GTPases [
19
]. In these formins, the FH1 and FH2 domains are flanked by
additional regulatory domains at the N-terminus, and by a Diaphanous-autoregulatory
domain (DAD) at the C-terminus. Binding of active Rho family proteins to the GTPase-
binding domain (GBD) triggers the activation of formins by disrupting the intramolecular
interaction between the DAD and the N-terminal Diaphanous-inhibitory domain (DID).
Several formins such as mDia2 [
20
,
21
] and the formin-like family members 2 (FMNL2) and
3 (FMNL3) [
22
–
24
] localize at lamellipodia and filopodia tips and have been implicated in
driving these protrusions.
The second group of proteins enhancing the rate and extent of actin filament elonga-
tion are Ena/VASP proteins. Vertebrates express Vasodilator-stimulated phosphoprotein
(VASP), mammalian Enabled (Mena), and Ena/VASP-like (Evl). All family members are
tetramers with a tripartite architecture harboring domains allowing for interactions with
FPPPP-containing receptors mediating subcellular positioning, actin monomers, profilin–
actin complexes and actin filaments [
10
]. In contrast to formins, single Ena/VASP tetramers
are only poorly processive and barely antagonize CP [
25
,
26
]. However, processivity and
resistance against CP increases dramatically upon oligomerization or clustering [
25
,
27
,
28
].
Previously, the clustering of VASP was thought to be mediated by the membrane deforming
and curvature sensing I-BAR (inverse Bar-domain) proteins such as IRSp53 (Insulin Re-
ceptor Substrate of 53 kDa) [
29
] or the scaffolding protein Lamellipodin (Lpd; also known
as Raph1) [
30
]. Very recently, however, it was shown that Ena/VASP clustering involves
Lpd but not I-BAR proteins and absolutely requires unconventional myosin-X (also known
as Myo10 or MyoX) [
31
]. Ena/VASP proteins localize to sites of active actin assembly in-
cluding focal adhesions (FA), SFs, the surface of bacterial pathogens and the tips of cellular
protrusions [
10
]. Consistent with their accumulation at the tips of filopodia, genetic removal
of all three Ena/VASP proteins was reported to severely perturb filopodia formation in
neuronal cells on poly-L-lysine, albeit the phenotype was less drastic on laminin [32,33].
Despite being composed of actin bundles and sharing common components, mi-
crospikes represent distinct molecular entities, as revealed by more recent work [
10
,
34
,
35
].
Microspikes, frequently capable of surfing along the plasma membrane, are an integral part
of the lamellipodium that mostly do not extend beyond the periphery of the membrane [
36
],
whereas filopodia, capable of protruding a few microns beyond the membrane [
37
], can
also form in the absence of lamellipodia and arise on the entire cell surface. Notably,
Ena/VASP-deficient B16-F1 cells virtually lack microspikes, but they can form numerous
filopodia upon treatment with the Arp2/3 complex inhibitor CK666 or ectopic expression
of MyoX and active mDia2 [
34
]. However, in spite of their ability to induce filopodia,
expression of either MyoX or active mDia2 fails to rescue microspike formation in these
mutants. In line with this notion, loss of MyoX completely abolishes microspikes in B16-F1
cells [
31
], whereas the knockdown of MyoX in endothelial cells decreases filopodia number
only by 50% [38].
The depletion of CP-ß in B16-F1 melanoma cells was previously shown to perturb
lamellipodia and cause an explosive formation of filopodia [
39
]. Since this phenotype was
very similar in Rat2 and NIH 3T3 fibroblasts, but was not observed in Ena/VASP-deficient
MV
D7
fibroblasts, it was hypothesized that Ena/VASP proteins are essential for filopodium
formation in the absence of CP [
39
]. To this end, in this study, we employed B16-F1 and NIH
3T3 cells and CRISPR/Cas9 technology followed by comprehensive analysis of filopodium
formation in derived mutant cells to experimentally test this hypothesis.

Cells 2023,12, 890 3 of 25
2. Materials and Methods
2.1. Constructs
Full-length Capzb encoding murine capping protein subunit beta (CapZ
β
) was ampli-
fied from an NIH 3T3 cDNA library and inserted into XhoI and EcoRI sites of plasmids
pEGFP-C1 and pEGFP-N1 (Clontech, Palo Alto, CA, USA). To allow for generation of stably
transfected cell lines, Capzb was also inserted into the XhoI and EcoRI sites of a pEGFP-C1
plasmid variant containing a Puromycin cassette (pEGFP-C1-Puro) [
34
]. The plasmid for
the expression of LifeAct fused to EGFP has been described [34,40].
For the generation of antigens, the coding sequence encompassing residues 1–527 of
murine FMNL2 was inserted into the BglII and SalI sites of pGEX 6P3 (GE Healthcare,
Munich, Germany). Accordingly, the sequence encoding residues 533–1028 of murine
FMNL3 was inserted into the BamHI and SalI sites of pGEX 6P1, and the sequence encoding
residues 94–473 of murine mDia2 was inserted into the EcoRI and SalI sites of pGEX 6P1.
All constructs were validated via sequencing.
2.2. Cell Culture and Transfection
NIH 3T3 fibroblasts (ATCC CRL-1658), B16-F1 mouse melanoma cells (ATCC CRL-
6323) and derived mutants were cultivated at 37
◦
C and 5% CO
2
in high-glucose DMEM cul-
ture medium (Lonza, Cologne, Germany) supplemented with 1% penicillin-streptomycin
(Biowest, Nuaille’, France), 10% FBS (Biowest) and 2 mM UltraGlutamine (Lonza). A total
of 3 h after seeding onto 35 mm diameter wells (Sarstedt, Nümbrecht, Germany), the cells
were transfected with 1
µ
g (B16-F1) or 3
µ
g (NIH 3T3) plasmid DNA using JetPRIME
transfection reagent (PolyPlus) at a ratio of 1
µ
g of DNA to 2
µ
L of the transfection reagent,
according to the manufacturer’s protocol. At 4 h post transfection, the transfection mixture
was replaced with fresh culture medium. Cells stably transfected with pEGFP-C1-Puro-
CapZ
β
were maintained with, additionally, 1.5
µ
g/mL puromycin. Used cell lines were
routinely authenticated following common guidelines by local authorities.
2.3. Genome Editing by CRISPR/Cas9
To generate sgRNAs of 20 nucleotides with high efficiency and minimal off-target
effects covering all possible splice variants, the DNA target sequence was pasted into a
CRISPR/Cas9 design tool (https://cctop.cos.uni-heidelberg.de/, accessed on 23 April
2020). For Capzb, the targeting sequence 5
0
-ACTGCGCCTTGGACCTGATG-3
0
was used
to target exon 2 of the gene. The sequence 5
0
-GTGATCCGAGCTCACATCTT-3
0
was used
to target exon 22 of the MYO10 gene. For the Fmnl2 and Fmnl3 genes, the sequences 5’-
TATGGGGAGGGTTCTTCACC-3
0
and 5
0
-TCTTGGACCCCAATGTAACA-3
0
, for targeting
exon 3 in the respective formins, were used. Each sequence was inserted into the BbsI
site of the pSpCas9(BB)-2A-Puro(PX459)V2.0 (Addgene plasmid ID: 62988) expression
plasmid [
41
]. Validation of the target sequences was performed by sequencing with a
5
0
-GGACTATCATATGCTTACCG-3
0
primer. At 24 h post transfection, cells were selected
in cell culture medium containing 2
µ
g/mL (for B16-F1-derived clones) or 3.5
µ
g/mL
(for NIH 3T3-derived clones) puromycin for 4 days and then cultivated for 24 h in the
absence of puromycin. For isolation of clonal knockout cell lines, single cells were seeded
by visual inspection into 96-well microtiter plates and expanded in conditioned culture
medium composed of used and sterile filtered medium and fresh medium at a ratio of 1:3.
Clones were verified on the genomic level using the TIDE sequence trace decomposition
web tool (https://tide.nki.nl/, accessed on 26 May 2020; [
42
]) and on the protein level by
immunoblotting using specific antibodies.
2.4. Protein Purification
GST-tagged mDia2 and FMNL2/3 were expressed in E. coli host Rosetta 2 (Sigma,
St. Louis, MO, USA) by induction with 1 mM isopropyl-
β
-D-thiogalactoside (Carl Roth,
Karlsruhe, Germany) at 24
◦
C for 12 h. The bacteria were harvested and lysed by ultra-
sonication in lysis buffer containing PBS, pH 7.4 supplemented with 2 mM DTT, 1 mM

Cells 2023,12, 890 4 of 25
EDTA, 5 mM benzamidine (Carl Roth), 0.1 mM AEBSF (AppliChem, Darmstadt, Germany),
Benzonase (1:1000, Merck, Darmstadt, Germany) and 5% (v/v) glycerol. The proteins
were then purified from bacterial extracts by affinity chromatography using glutathione-
conjugated agarose (Macherey-Nagel, Düren, Germany) and eluted from the column with
20 mM reduced glutathione (Carl Roth) in PBS supplemented with 2 mM DTT, 1 mM EDTA,
5 mM benzamidine, 0.1 mM AEBSF and 5% (v/v) glycerol using standard procedures. The
GST tag was cleaved off by PreScission protease (GE Healthcare), and the GST tag was
then absorbed on fresh glutathione-conjugated agarose. The proteins in the flow through
were further purified by size-exclusion chromatography (SEC) using a HiLoad 26/600
Superdex 200 column (GE Healthcare) controlled by an Äkta Purifier System. The fractions
containing mDia2, FMNL2 and
−
3 were pooled and dialyzed against immunization buffer
(150 mM NaCl, 25 mM Tris/HCl, pH 8.0) for generation of polyclonal antibodies.
2.5. Antibodies
Polyclonal antibodies against recombinant fragments of mDia2, FMNL2 and FMNL3
were raised by immunization of female New Zealand white rabbits. This was followed by
antigen-affinity purification against the same antigens coupled to CNBr-activated sepharose
4B (GE Healthcare) according to the manufacturer’s protocol. Briefly, after passing the sera
over the column, the resin was washed with 300 mL of PBS, pH 7.4 and bound antibodies
were eluted with 0.1 M acetic acid, pH 2.3. The fractions containing the antibodies were
pooled, and the pH was immediately adjusted to 7.5 using 1 M Tris/HCl, pH 8.5 followed
by dialysis against PBS buffer supplemented with 55% (v/v) glycerol for long-term storage
at
−
20
◦
C. The immunization of rabbits for the generation of polyclonal antibodies was
conducted in accordance with national guidelines for the care and maintenance of laboratory
animals and approved by the Hannover Medical School Institutional Animal Care Facility
and the Lower Saxony State Office for Consumer Protection and Food Safety (LAVES) under
the application number 18A255 to J.F. For immunoblotting, polyclonal antibodies against
FMNL2 (1:1000 dilution), FMNL3 (1:1000, dilution) and MyoX (1:1000 dilution, [
31
]) and the
monoclonal antibodies directed against the capping protein
α
1/
α
2 subunits mAb B5 12.3 (1:4
hybridoma supernatant, Developmental Hybridoma Bank (deposited by J. Cooper), Iowa
City, IA, USA), the capping protein
β
2 subunit mAb 3F2.3 (1:4 hybridoma supernatant, De-
velopmental Hybridoma Bank (deposited by J. Cooper)), pan anti-actin (1:1000, # ab119952,
Abcam, Boston, MA, USA) and GAPDH (1:50,000; #CB1001-500UG, Merck, Darmstadt,
Germany) were used. Primary antibodies in immunoblots were either visualized using
phosphatase-coupled anti-rabbit (1:1000, #115-055-114, Dianova, Hamburg, Germany) and
anti-mouse (1:1000; #115-055-62, Dianova) antibodies or by enhanced chemiluminescence
using peroxidase-coupled anti-mouse IgG (Dianova; #115-035-062; 1:10,000 dilution).
For immunofluorescence, the following primary antibodies were used: rabbit anti-
VASP (1:1000 dilution, [
34
]), rabbit anti-FMNL2 (1:1000 dilution), rabbit anti-FMNL3 (1:1000,
dilution), rabbit anti-MyoX (1:1000 dilution [
31
]), rabbit anti-WAVE2 (1:1000 dilution [
31
]
and the monoclonal anti-vinculin antibody hVIN-1 (1:1000 dilution; #V9131, Sigma). Pri-
mary antibodies were visualized in immunocytochemistry with Alexa-488-conjugated
goat-anti-rabbit (1:1000 dilution; #A-11034, Invitrogen) or goat-anti-mouse antibodies
(1:1000 dilution; #A-11029, Invitrogen). To enhance EGFP signals, Alexa488-conjugated
anti-EGFP nanobodies (1:1000 dilution, [43]) were used.
2.6. Immunoblotting
For the preparation of whole-cell lysates, cells were cultivated to 80–100% confluency
and trypsinized. Cell pellets were washed twice with cold PBS and lysed with 400
µ
L of cold
RIPA buffer (150 mM NaCl, 1.0% Triton-X-100, 0.5% sodium deoxycholate, 0.1% sodium
dodecyl sulfate (SDS), 50 mM Tris, pH 8.0), then supplemented with 5 mM benzamidine
(Carl Roth), 0.1 mM AEBSF (AppliChem, Darmstadt, Germany) and benzonase (1:1000,
Merck, Darmstadt, Germany) for 25 min at 4
◦
C on a wheel rotator. Subsequently, the SDS
concentration was adjusted to 0.3% final concentration, and cells were further lysed for

Cells 2023,12, 890 5 of 25
20 min at 4
◦
C on a wheel rotator. Then, cell lysate was centrifuged at 4
◦
C for 5 min at
20,000
×
g, and the supernatant was resuspended in an appropriate volume of 3
×
SDS
sample buffer (150 mM Tris/HCl, pH 6.8, 30% glycerol, 3% SDS, 3% β-mercaptoethanol).
Total proteins of cell lysates were resolved by SDS-PAGE, transferred by semi-dry
blotting using a Trans-Blot SD Semi-Dry Transfer Cell (Bio Rad, Feldkirchen, Germany)
at 16 V onto nitrocellulose membranes (Hypermol, Hannover, Germany) and blocked
with NCP buffer (10 mM Tris/HCl pH 8.0, 150 mM NaCl, 0.05% Tween-20, 0.02% NaN
3
)
containing 4% bovine serum albumin (BSA) for at least 30 min. Primary antibodies were
incubated overnight in NCP buffer. After washing of the membranes five times with NCP
buffer and incubation with secondary, phosphatase-conjugated antibodies for at least 4 h,
the blots were developed with 20 mg/mL 5-brom-4-chlor-3-indolylphosphate-p-toluidin
(BCIP) in 0.1 M NaHCO
3
, pH 10.0. For quantification of proteins, the immunoblots were
developed by enhanced chemiluminescence, and analysis was performed with ImageJ [
44
].
Uncropped scans of immunoblots and gels are shown in Figure S11.
2.7. Immunofluorescence and Imaging
For immunofluorescence labeling, cells were fixed for 20 min in pre-warmed PBS, pH
7.3 containing 4% PFA and 0.06% picric acid and, subsequently, were washed three times
with PBS supplemented with 100 mM glycine. The cells were then permeabilized with 0.1%
Triton X-100 in PBS for 35 s and blocked with PBG (PBS, 0.045% cold fish gelatin and 0.5%
BSA). Primary antibodies were incubated overnight, followed by washing of the specimens
five times with PBG and incubation with respective secondary antibodies for at least 2 h. F-
actin was visualized with Atto550-phalloidin (1:250 dilution, #AD 550–82, Atto-Tec, Siegen,
Germany), and DNA was visualized with 4
0
,6-diamidino-2-phenylindole (DAPI) (1:1000
dilution, Sigma). The 16-bit images of fixed cells were captured with an Olympus XI-81
inverted microscope equipped with an UPlan FI 100
×
/1.30 NA oil immersion objective or
a Zeiss LSM 980 with Airyscan 2 equipped with a Plan-Apochromat 63
×
/1.4 NA oil DIC
objective using 405 nm, 488 nm and 594 nm laser lines. Emitted light was detected in the
wavelength ranges of 410–473 nm, 490–561 nm and 576–695 nm, respectively.
Time-lapse imaging of live cells was performed using an Olympus XI-81 inverted micro-
scope (Olympus, Hamburg, Germany) driven by Metamorph software (Molecular Devices,
San Jose, CA, USA) and equipped with objectives specified below and a CoolSnap EZ camera
(Photometrics, Tucson, AZ, USA). Cells were seeded onto 35 mm glass bottom dishes (Ibidi,
Planegg-Martinsried, Germany), coated for 1 h with either 25 mg/mL laminin (Sigma), in
the case of B16-F1 cells and derived clones, or with 20
µ
g/mL fibronectin (Roche, Penzberg,
Germany), in the case of NIH 3T3 fibroblasts and their derivatives, maintained in imaging
medium composed of F-12 Ham Nutrient Mixture with 25 mM HEPES (Sigma) to compensate
for the lack of CO
2
and supplemented with 10% FBS (Biowest), 1% Penicillin-Streptomycin
(Biowest), 2 mM stable L-glutamine (Biowest) and 2.7 g/L D-glucose (Carl Roth, Karlsruhe,
Germany) in an Ibidi Heating System at 37
◦
C. For 2D random motility assays, B16-F1 cells
were seeded at low density (~5
×
10
4
cells/mL) onto the dishes and allowed to adhere for
3 h. Subsequently, the medium was exchanged with imaging medium, the chamber was
mounted into a heating system and cells were recorded by time-lapse, phase-contrast imaging
with 16-bit image depth at 60 s intervals for 3 h using an UPlan FL N 4
×
/0.13 NA objective
(Olympus) with additional 1.6
×
optovar magnification. NIH 3T3 and derived cells were
allowed to adhere for 3 h after seeding and were imaged at 10 min intervals for 10 h using
an Uplan FL N 4
×
/0.13 NA objective (Olympus, Hamburg, Germany). Single-cell tracking
was performed with MTrackJ in ImageJ [
44
,
45
]. Analyses of cell speed and cell trajectories,
turning angles and mean square displacements were performed in Excel (Microsoft, Redmond,
WA, USA) using a customized macro [
46
]. Cells that contacted each other or divided were
excluded from analysis. The directionality index was calculated by dividing the shortest
distance between starting and end points (d) by the actual cell trajectories (D). Behavior of
migrating cells on laminin at high magnification was recorded by time-lapse imaging at 5 s
intervals for 15 min using an UPlan FI 100×/1.30 NA oil immersion objective (Olympus).

Cells 2023,12, 890 6 of 25
2.8. Quantification of F-Actin Content by Flow Cytometry
Flow cytometry was performed with an FACSAria III Fusion flow cytometer (Becton
Dickinson, Franklin Lake, NJ, USA) driven by FACSDiva Vers. 8.0.1. software. For
fluorescence-activated cell sorting of reconstituted CP-KO cells stably expressing EGFP-
CapZ
β
2, EGFP was excited with a laser at 488 nm, and emission at 530 nm was detected
with a band-pass filter. EGFP-positive cells were collected into a sterile 15 mL screw-cap
tube containing cell culture medium with 1.5
µ
g/mL puromycin for further cultivation.
To quantify cellular F-actin content, the cells were trypsinized, fixed and permeabilized in
solution and then counted, and approximately 2
×
10
5
cells were stained for 2 h with 3
µ
M
Atto550-conjugated phalloidin (Attotec, Siegen, Germany) in 400
µ
L PBS. After washing
five times with PBS, the cells were analyzed by flow cytometry using 561 nm laser light
excitation. Emission at 586 nm was detected using a band-pass filter. The number of events
counted was approximately 50,000 for each cell line in each run. Further analysis was
performed with FlowJo software (Becton Dickinson). Data are presented as the x-fold
change in phalloidin intensity in comparison to B61-F1 control.
2.9. Quantification of the F- to G-Actin Ratio and of Global Actin Levels
To determine the F- to G-actin ratio in B16-F1 wild-type and CP-deficient cells, nearly
confluent cultures, corresponding to about 4–5
×
10
6
cells, were first rinsed twice with
ice-cold PBS. Cells from each plate were directly detached from the plate with a cell scraper
using 300
µ
L of cold lysis buffer containing 20 mM Hepes, pH 7.2, 100 mM NaCl, 10 mM
KCl, 2 mM MgCl
2
, 5 mM KPO
4
, 5 mM EGTA, 5 mM ATP, 2 mM DTT, 5% sucrose, 0.5%
Triton X-100, 0.5% NP-40, 5 mM benzamidine, 0.1 mM AEBSF and 0.75
µ
M phalloidin
(Sigma). The crude lysates were then transferred into 1.5 mL reaction tubes and placed
on a rotary shaker for further homogenization at 4 ◦C. After 30 min, 200 µL samples were
centrifuged at 150,000
×
gfor 1 h using an Optima tabletop ultracentrifuge (Beckman
Instruments, Palo Alto, CA, USA). Subsequently, the samples of the supernatant and
the pellet fractions were brought to 300
µ
L with SDS sample buffer, and aliquots were
subjected to SDS-PAGE followed by Coomassie Blue staining or immunoblotting. The blots
were probed with pan anti-actin antibody at a 1:1000 dilution overnight. Primary actin
antibodies in these immunoblots were visualized by enhanced chemiluminescence using
the ChemiDoc MP Imaging System driven by Image Lab software (Biorad, Hercules, CA,
USA). After background subtraction of 16-bit images, the amount of actin in the pellet
and supernatant fractions was determined densitometrically using ImageJ software. For
quantification of global actin levels, proteins in total cellular lysates were separated by
SDS-PAGE and blotted on nitrocellulose, and blots were incubated with anti-actin and
anti-GAPDH antibodies. After background subtraction of 16-bit images, band intensities
of actin were normalized to respective GAPDH signals, and relative protein levels were
calculated in CP-KO mutants compared with the B16-F1 control.
2.10. Quantification of Peripheral Filopodia
Brightest point projection of 3D reconstructions from confocal Airyscan sections were
used to visualize filopodia of phalloidin-stained cells at high resolution, regardless of
their morphology. Filopodium length was defined as the distance of the filopodium shaft
extending beyond the periphery of the plasma membrane and its distal tip. The length and
the number of peripheral filopodia per cell were quantified manually with ImageJ.
2.11. Analysis of Focal Adhesions
FA parameters of NIH 3T3 fibroblasts and derived mutants were inferred from confo-
cal images of vinculin-stained cells using a customized ImageJ macro to facilitate analysis.
Semi-automatic processing of 16-bit images recorded at identical settings included gradual
background subtractions using a rolling ball radius of 50 pixels to obtain pre-processed images
for subsequent analysis of intensity profiles. For segmentation of FAs, the resulting images
were further processed by two additional background subtraction steps using rolling ball

Cells 2023,12, 890 7 of 25
radii of 15 pixels followed by binarization of obtained images using the Otsu thresholding
method [
47
]. The regions obtained were redirected to the pre-processed images, which were
then analyzed with the analyze particles plugin in ImageJ using a minimum particle size of
0.25
µ
m
2
. This yielded binary masks of FAs for visual display and specific FA parameters
such as number, size, length and intensity, which were further analyzed using Microsoft Excel.
2.12. Statistical Analyses
Quantitative experiments were performed at least in triplicates to minimize environ-
mental bias or unintentional error. Raw data were processed in Excel. Statistical analyses
were performed with Origin 2021 (OriginLab Corporation, Northampton, MA, USA). All
datasets were tested for normality by the Shapiro–Wilk test. Statistical differences between
normally distributed datasets of two groups were determined by t-test, and non-normally
distributed datasets of two groups were determined by a non-parametric Mann–Whitney U
rank sum test. For comparison of more than two groups, statistical significance of normally
distributed data was examined by one-way ANOVA and a Tukey Multiple Comparison
test. In the case of non-normally distributed data, the non-parametric Kruskal–Wallis test
and Dunn’s Multiple Comparison test were used. Statistical differences were defined as
*p
≤
0.05, ** p
≤
0.01, *** p
≤
0.001 as well as n.s., not significant, and are displayed and
stated in figures and figure legends, respectively.
3. Results
3.1. Loss of CP in B16-F1 Cells Triggers the Massive Formation of Filopodia and Impairs
2D-Cell Migration
To test the hypothesis of whether Ena/VASP proteins are essential for filopodia formation
in CP-deficient cells [
39
], we first disrupted the single gene (CapZb) encoding the ß-subunit of
CP in B16-F1 mouse melanoma cells using CRISPR/Cas9 technology. Respective protein loss
in independent clonal cell lines was validated by TIDE sequence trace decomposition analyses
of amplified genomic target sites (Figure S1A) [
42
] and, finally, confirmed by sequencing of
the target site and immunoblotting (Figure 1A). Given that CP operates only as a functional
heterodimer, consistent with previous work [
39
,
48
], loss of the ß-subunit also resulted in a
markedly reduced expression level of the
α
-subunit. We then analyzed actin filament (F-actin)
distribution in B16-F1 and CP-KO cells after phalloidin staining. As opposed to B16-F1 control,
where almost 50% of the cells displayed prominent smooth lamellipodia with numerous
microspikes, approximately 12–14% of the CP-KO cells developed strongly compromised
lamellipodia with bulbous, exceptionally strong ruffling cell fronts with many protruding
filopodia (Figures 1B and S2). The remaining 86–88% of the mutant cells were apparently not
polarized and also exhibited massive formation of highly dynamic filopodia around the entire
cell periphery and on the dorsal surface (Movies S1–S3). To ensure that the observed mutant
phenotypes were CP-specific, we then analyzed reconstituted CP-KO mutant cells ectopically
expressing the ß2-subunit of CP fused to EGFP. Re-expression of CP-ß2 not only restored
accumulation of the endogenous CP-
α
-subunit (Figure 1C) but also rescued cell morphology
and F-actin distribution (Figure 1D).
Since 2D migration on flat surfaces is primarily driven by actin assembly in the lamel-
lipodium, we then used phase-contrast, time-lapse imaging to measure rates of cell migra-
tion in parental B16-F1 cells and two independent CP-KO cell lines on laminin (
Movie S4
).
As opposed to control cells, which migrated at 1.47
±
0.38
µ
m/min (
mean ±SD
), cell
speed was reduced by about 70% in both CP-deficient mutant cell lines (clones #9 and
#22) to
0.40 ±0.25 µ
m/min and 0.54
±
0.28
µ
m/min, respectively (Figure 1E). This was
accompanied by increases in directionality in both CP-KO mutants, making them 55% or
32% more directional compared to the wild type (Figure 1F). Finally, we calculated the
mean square displacement (MSD) in wild-type and mutant cells to assess their effective
directional movement. Despite their higher directionality, presumably owing to their
markedly slower motility, both CP-deficient cell lines displayed drastically lower MSD
values as compared to the B16-F1 control (Figure 1G). Taken together, these results are

Cells 2023,12, 890 8 of 25
consistent with previous work [
39
,
48
–
50
] showing that CP is critical for effective 2D-cell
migration, even though complete loss of CP, as shown here, caused more severe motility
defects as compared to those previously reported for CP-depleted cells [49,50].
Cells 2023, 12, x FOR PEER REVIEW 8 of 27
the dorsal surface (Movies S1–S3). To ensure that the observed mutant phenotypes were
CP-specific, we then analyzed reconstituted CP-KO mutant cells ectopically expressing
the ß2-subunit of CP fused to EGFP. Re-expression of CP-ß2 not only restored accumula-
tion of the endogenous CP-α-subunit (Figure 1C) but also rescued cell morphology and
F-actin distribution (Figure 1D).
Since 2D migration on flat surfaces is primarily driven by actin assembly in the la-
mellipodium, we then used phase-contrast, time-lapse imaging to measure rates of cell
migration in parental B16-F1 cells and two independent CP-KO cell lines on laminin
(Movie S4). As opposed to control cells, which migrated at 1.47 ± 0.38 µm/min (mean ±
SD), cell speed was reduced by about 70% in both CP-deficient mutant cell lines (clones
#9 and #22) to 0.40 ± 0.25 µm/min and 0.54 ± 0.28 µm/min, respectively (Figure 1E). This
was accompanied by increases in directionality in both CP-KO mutants, making them 55%
or 32% more directional compared to the wild type (Figure 1F). Finally, we calculated the
mean square displacement (MSD) in wild-type and mutant cells to assess their effective
directional movement. Despite their higher directionality, presumably owing to their
markedly slower motility, both CP-deficient cell lines displayed drastically lower MSD
values as compared to the B16-F1 control (Figure 1G). Taken together, these results are
consistent with previous work [39,48–50] showing that CP is critical for effective 2D-cell
migration, even though complete loss of CP, as shown here, caused more severe motility
defects as compared to those previously reported for CP-depleted cells [49,50].
Figure 1.
Loss of CP in B16-F1 cells triggers the massive formation of filopodia and impairs 2D-cell
migration. (
A
) Immunoblot confirming the elimination of CP-ß in two independent single-knockout
B16-F1 mutants (clones #9 and #22). The loss of the ß-subunit also led to the almost complete loss of
the
α
-subunit. Loading control: GAPDH. (
B
) Morphology of representative B16-F1 cells with a smooth
leading edge and embedded microspikes (left) and of CP-KO mutants that were either polarized and
exhibited strong ruffling of the leading edge (middle) or were unpolarized (right). Cells migrating on
laminin were stained for F-actin with phalloidin. Note the dramatically increased formation of filopodia
in the mutant cells. (
C
,
D
) Reconstitution of CP-KO cells with EGFP-tagged CP-ß restores expression of
the CP-
α
-subunit (
C
) and rescues the phenotype (
D
). (
E
) Loss of CP in B16-F1 cells decreases random
2D-cell migration on laminin. At least three time-lapse movies from three independent experiments
were analyzed for each cell line. The boxes in box plots indicate 50% (25–75%) and whiskers (5–95%)
of all measurements, with dashed black lines depicting the medians and arithmetic means highlighted
in blue. (
F
) Directionality increased upon inactivation of CP. Bars represent arithmetic means
±
SD.
(
G
) Analyses of mean square displacement of wild-type versus mutant cells. Error bars represent means
±
SEM. (
E
,
F
) Non-parametric, Kruskal–Wallis test and Dunn’s Multiple Comparison test were used to
reveal statistically significant differences between datasets. * p
≤
0.05, *** p
≤
0.001; n.s.: not significant.
n: number of cells analyzed from at least three independent experiments.

Cells 2023,12, 890 9 of 25
3.2. Loss of CP in B16-F1 Cells Increases Global Levels of Filamentous and Total Actin
In agreement with previous work analyzing CP-depleted Dictyostelium or B16-F10
mouse melanoma cells [
49
,
50
], elimination of CP in B16-F1 cells caused a significant in-
crease in F-actin intensity, as evidenced by epifluorescence imaging of phalloidin-stained
wild-type and mutant cells at identical settings (Figure 2A). To allow the most accurate
quantification of F-actin content in CP-deficient B16-F1 cells, we trypsinized adherent wild-
type and mutant cells and fixed and stained them with fluorescent phalloidin at saturating
conditions in solution. The labeled specimens were then analyzed by flow cytometry, which
is a powerful tool for the precise quantitation of fluorescence intensities on the single-cell
level [
51
]. As expected, both CP-KO mutants exhibited markedly increased phalloidin
fluorescence intensities as compared to the B16-F1 control (Figure 2B). Quantification of
relative fluorescence intensities revealed that the F-actin content was increased 4- to 5-fold
in the CP-KO cells compared with the B16-F1 control (Figure 2C), supporting the notion
that elimination of CP elicits unleashed actin assembly. Notably, re-expression of EGFP-
labeled CP-ß in CP-KO cells almost fully reverted phalloidin intensity in reconstituted
cells to wild-type levels (Figure 2D). Given the drastically increased F-actin levels in the
mutants, we then asked how loss of CP would affect the F- to G-actin ratio in the CP-KO
cells. The amounts of free globular G-actin and filamentous F-actin were determined by
differential precipitation of F-actin (pellet fraction) and G-actin (supernatant fraction) in to-
tal cell lysates after ultracentrifugation. After SDS-PAGE and Coomassie Blue staining, we
noticed a prominent band at 42 kDa in the pellet fractions of the mutant cells, presumably
representing actin (Figure 2E). This was confirmed by immunoblotting with an anti-actin
antibody. Densitometric analysis and quantification of respective actin signals revealed
a dramatically increased F- to G-actin ratio of more than 7:1 in both CP-KO mutants, as
opposed to a ratio of 1.2:1 in the B16-F1 control (Figure 2E). Given the critical function of
CP in maintaining the available pool of polymerization-competent monomeric actin, finally,
we analyzed, by quantitative immunoblotting, whether the markedly increased F-actin
content in CP-KO cells also altered global actin expression. Densitometry revealed that total
actin levels, normalized to GAPDH expression, were approximately 2.8-fold higher in the
KO mutants compared to the B16-F1 control (Figure S3). Taken together, these data suggest
that, due to excessive actin polymerization, CP-deficient cells apparently compensate for
depletion of the G-actin pool by stronger expression of actin.
3.3. Loss of CP in NIH 3T3 Fibroblasts Induces the Massive Formation FAs and SFs Instead of
Filopodia on Fibronectin
To corroborate the filopodial and migratory phenotypes after loss of CP in a more
strongly adherent cell type, we then disrupted the ß-subunit of CP by CRISPR/Cas9 in
NIH 3T3 fibroblasts. Respective loss of CP-ß in independent clonal cell lines was again
validated by TIDE analysis of amplified genomic target sites and, finally, confirmed by
immunoblotting (Figures 3A and S1B). Comparable to CP-deficient B16-F1 cells, in NIH 3T3
cells, loss of the ß-subunit of CP also resulted in markedly reduced expression or stability
of the
α
-subunit (Figure 3A). We then examined F-actin distribution in parental NIH 3T3
cells and derived CP-KO cells after phalloidin staining. However, in variance to previous
work [
39
], and as opposed to CP-deficient B16-F1 cells, CP-deficient NIH 3T3 cells did
not form numerous filopodia but instead developed a conspicuously dense meshwork of
SFs that almost completely filled the mutant cells (Figure 3B), suggesting that CP-KO cells
become extraordinarily adhesive [
52
]. To experimentally test this hypothesis and examine
the consequences on fibroblast adhesion after loss of CP, wild-type and CP-deficient NIH
3T3 cells migrating on fibronectin were labeled for the focal adhesion (FA) marker protein
vinculin and assessed for various features (Figure 3C). To this end, images captured at
identical settings were processed into binary images using a customized macro, allowing
global and unbiased assessment of multiple FA parameters. Notably, vinculin intensity was
markedly increased in the CP-deficient mutants by 54.1
±
36.1% (clone #4) and 42.2
±
33.4%
(clone #9), as compared to the parental NIH 3T3 cell line (Figure 3D). Moreover, the number

Cells 2023,12, 890 10 of 25
of FAs in CP-KO cells was also increased by more than 50%, to 302
±
128 and 287
±
104, as
compared to the wild type, with 138
±
43 (Figure 3E). Quantification of FA size, furthermore,
revealed an increase in CP-KO cells by more than 50%, to 1.7
±
0.4
µ
m
2
and 1.7
±
0.4
µ
m
2
,
as compared to NIH 3T3 control cells, with 1.1
±
0.3
µ
m
2
(Figure 3F). In addition, FAs in the
mutant cells were considerably longer and wider as compared to control (Figure S4). Thus,
loss of CP in NIH 3T3 fibroblasts apparently appears to dramatically improve adhesion to
their preferred cell substrate fibronectin but does not lead to excessive filopodia formation.
Cells 2023, 12, x FOR PEER REVIEW 10 of 27
Figure 2. Loss of CP results in markedly increased F-actin levels. (A) Representative examples of
B16-F1 cells and derived CP-KO mutant stained with phalloidin for the F-actin cytoskeleton and
imaged at identical settings. Note the much brighter phalloidin signal in CP-KO cells. (B) Flow cy-
tometry of phalloidin-stained B16-F1 cells and two independent CP-KO mutant cell lines. (C) Quan-
tification of F-actin content from flow cytometry experiments shown in (B). (D) Flow cytometry of
phalloidin-stained B16-F1 wild-type, CP-KO and reconstituted CP-KO cells expressing EGFP-CP-β.
(E) Immunoblot depicting actin levels in pellet (P) and supernatant (S) fractions of B16-F1 and de-
rived CP-KO mutants. The corresponding Coomassie Blue-stained gel is shown above. Note the
prominent band of approximately 42 kDa in the pellet fractions of the mutant cells, which most
likely represents actin. (F) Quantification of actin in pellet (P) and supernatant (S) fractions from
immunoblots shown in (E). (C,F) Bars represent arithmetic means ± SD. Non-parametric, Kruskal–
Wallis test and Dunn’s Multiple Comparison test (C) and one-way ANOVA and Tukey Multiple
Comparison test (F) were used to reveal statistically significant differences between datasets. ** p ≤
0.01, *** p ≤ 0.001; n.s.: not significant. n: number of independent experiments using approximately
5 × 104 cells for each cell line (B,D) or the number of independent experiments (C,F).
3.3. Loss of CP in NIH 3T3 Fibroblasts Induces the Massive Formation FAs and SFs Instead of
Filopodia on Fibronectin
To corroborate the filopodial and migratory phenotypes after loss of CP in a more
strongly adherent cell type, we then disrupted the ß-subunit of CP by CRISPR/Cas9 in
NIH 3T3 fibroblasts. Respective loss of CP-ß in independent clonal cell lines was again
validated by TIDE analysis of amplified genomic target sites and, finally, confirmed by
immunoblotting (Figures 3A and S1B). Comparable to CP-deficient B16-F1 cells, in NIH
Figure 2.
Loss of CP results in markedly increased F-actin levels. (
A
) Representative examples
of B16-F1 cells and derived CP-KO mutant stained with phalloidin for the F-actin cytoskele-
ton and imaged at identical settings. Note the much brighter phalloidin signal in CP-KO cells.
(
B
) Flow cytometry of phalloidin-stained B16-F1 cells and two independent CP-KO mutant cell lines.
(
C
) Quantification of F-actin content from flow cytometry experiments shown in (
B
). (
D
) Flow cy-
tometry of phalloidin-stained B16-F1 wild-type, CP-KO and reconstituted CP-KO cells expressing
EGFP-CP-
β
. (
E
) Immunoblot depicting actin levels in pellet (P) and supernatant (S) fractions of
B16-F1 and derived CP-KO mutants. The corresponding Coomassie Blue-stained gel is shown above.
Note the prominent band of approximately 42 kDa in the pellet fractions of the mutant cells, which
most likely represents actin. (
F
) Quantification of actin in pellet (P) and supernatant (S) fractions
from immunoblots shown in (
E
). (
C
,
F
) Bars represent arithmetic means
±
SD. Non-parametric,
Kruskal–Wallis test and Dunn’s Multiple Comparison test (
C
) and one-way ANOVA and Tukey
Multiple Comparison test (
F
) were used to reveal statistically significant differences between datasets.
** p
≤
0.01, *** p
≤
0.001; n.s.: not significant. n: number of independent experiments using approxi-
mately 5 ×104cells for each cell line (B,D) or the number of independent experiments (C,F).

Cells 2023,12, 890 11 of 25
Cells 2023, 12, x FOR PEER REVIEW 12 of 27
Figure 3. Loss of CP in NIH 3T3 fibroblasts amplifies SF and FA formation and impairs 2D-cell
migration. (A) Immunoblot confirming the elimination of CPß in two independent single-knockout
NIH 3T3 mutants. The loss of the CPß-subunit also led to the almost complete loss of the CPα-
subunit. Loading control: GAPDH. (B) Morphologies of representative NIH 3T3 and CP-KO mutant
cells migrating on fibronectin and stained for F-actin with phalloidin. Note the dramatically in-
creased formation of SFs in the mutant cells. (C) Representative micrographs of NIH 3T3 and a de-
rived CP-KO mutant cell displaying vinculin staining before (upper panel) and after processing
with a customized Fiji macro (lower panel). (D) Quantification of vinculin intensities in FAs. (E)
Quantification of FA number. (F) Quantification of FA area. (G) Loss of CP results in decreased cell
speed. (H) Analyses of mean square displacement of wild-type versus mutant cells. Respective sym-
bols and error bars represent means ± SEM. (D–G) The boxes in box plots indicate 50% (25–75%)
and whiskers (5–95%) of all measurements, with dashed black lines depicting the medians and arith-
metic means highlighted in blue. Non-parametric Kruskal–Wallis test and Dunn’s Multiple Com-
parison test were used to reveal statistically significant differences between datasets. *** p ≤ 0.001;
n.s.: not significant. n: number of cells analyzed from at least three independent experiments.
Figure 3.
Loss of CP in NIH 3T3 fibroblasts amplifies SF and FA formation and impairs 2D-cell
migration. (A) Immunoblot confirming the elimination of CPß in two independent single-knockout
NIH 3T3 mutants. The loss of the CPß-subunit also led to the almost complete loss of the CP
α
-subunit.
Loading control: GAPDH. (
B
) Morphologies of representative NIH 3T3 and CP-KO mutant cells
migrating on fibronectin and stained for F-actin with phalloidin. Note the dramatically increased
formation of SFs in the mutant cells. (
C
) Representative micrographs of NIH 3T3 and a derived CP-KO
mutant cell displaying vinculin staining before (upper panel) and after processing with a customized
Fiji macro (lower panel). (
D
) Quantification of vinculin intensities in FAs. (
E
) Quantification of FA
number. (
F
) Quantification of FA area. (
G
) Loss of CP results in decreased cell speed. (
H
) Analyses
of mean square displacement of wild-type versus mutant cells. Respective symbols and error bars
represent means
±
SEM. (
D
–
G
) The boxes in box plots indicate 50% (25–75%) and whiskers (5–95%)
of all measurements, with dashed black lines depicting the medians and arithmetic means highlighted
in blue. Non-parametric Kruskal–Wallis test and Dunn’s Multiple Comparison test were used to
reveal statistically significant differences between datasets. *** p
≤
0.001; n.s.: not significant. n:
number of cells analyzed from at least three independent experiments.

Cells 2023,12, 890 12 of 25
Finally, we assessed the consequences of CP deficiency on fibroblast 2D-cell migra-
tion on fibronectin by phase-contrast, time-lapse imaging of NIH 3T3 and CP-KO cells
(Figure 3G and Movie S5). In both CP-KO mutants, cell speed (0.10
±
0.06
µ
m/min
and 0.11
±
0.06
µ
m/min) was significantly reduced as compared to NIH 3T3 control
(
0.29 ±0.08 µm/min
). Consistently, the CP-KO mutants exhibited much lower MSD val-
ues when compared to the NIH 3T3 wild type (Figure 3H). Thus, despite their highly
different phenotypes regarding filopodia formation, both B16-F1- and NIH 3T3-derived
mutants exhibited strongly compromised 2D-cell migration after loss of CP.
3.4. CP-Deficent NIH 3T3 Cells form Numerous Filopodia on Poorly Adhesive Substrates
The strongly amplified formation of integrin-based cell adhesions, as well as the exces-
sive spreading of NIH 3T3-derived CP-KO mutants on fibronectin, suggested enhanced
contractility of FA-anchored SFs, likely resulting in cells with highly stretched membranes.
Interestingly, it has recently been shown that actin-based protrusions such as lamellipo-
dia adapt to changes in membrane tension imposed by branched actin networks, such
that large lamellipodia are formed at low tension, whereas their formation is perturbed
at high tension [
53
]. Notably, we found that lamellipodia formation in NIH 3T3 fibrob-
lasts was suppressed after loss of CP, as evidenced by immunostaining the cells with the
WRC component WAVE2 (Figure S5). Thus, in addition to the relevance of CP for lamel-
lipodium formation, this could also mean that filopodia formation in NIH 3T3-derived
CP-KO cells on fibronectin is suppressed because the force generated by nascent filopodia
is not sufficient to overcome the high tension of the membrane. Since it is not possible to
distinguish between these two possibilities without elaborate biophysical methods such
as atomic force microscopy (AFM) or the use of novel tension sensors probes reporting
membrane tension changes through their fluorescence lifetime [
54
], we therefore asked
whether filopodium formation could be induced by seeding the cells on poorly adhesive
substrates. As shown in Figure 4, CP-KO cells seeded on poly-L-lysine or directly on glass
adhered poorly but formed numerous filopodia, comparable to CP-deficient B16-F1 cells
plated on laminin. Thus, the striking filopodia phenotype upon loss of CP appears to be
cell-type- and context-dependent.
3.5. Loss of CP in Ena/VASP-Deficient B16-F1 Cells Does Not Abrogate Filopodium Formation
In previous work, it has been hypothesized that Ena/VASP proteins are obligatory for
filopodium formation in the absence of CP [
39
]. To confirm or challenge this hypothesis,
we first examined VASP localization in CP-deficient B16-F1 cells. CP-KO cells stained
for the F-actin cytoskeleton and endogenous VASP indeed showed conspicuous VASP
clusters at the distal tips of filopodia (Figure 5A). Then, we disrupted the ß-subunit of CP
by CRISPR/Cas9 in Ena/VASP/Mena-deficient B16-F1 cells (EVM-KO), recently shown
to virtually lack microspikes but still be capable of forming filopodia upon transient
expression of active mDia2 and unconventional myosin-X or by pharmacological inhibition
of the Arp2/3 complex [
34
]. Loss of CP-ß in independent clonal cell lines, denoted as
EVM/CP-KO cells, was again validated by TIDE analysis of amplified genomic target sites
and confirmed by immunoblotting (Figures 5B and S1A). Both independent EVM/CP-
KO mutant cells (clones #4 and #14) were highly similar, but, to our surprise, they still
formed numerous filopodia, similar to the CP-KO cells, as determined by confocal Airyscan
imaging of cells stained with phalloidin migrating on laminin (Figure 5C and Movie S6).
Quantification further revealed filopodia to be about 20% shorter in EVM/CP-KO mutants
(2.2
±
0.5 and 2.2
±
0.5
µ
m) as compared to CP-KO control cells (2.8
±
0.8 and 2.8
±
0.7
µ
m)
(Figure 5D). Notably, the quantification of the number of peripheral filopodia, on the other
hand, showed no noticeable differences between these cell lines (Figure 5E), suggesting
that Ena/VASP proteins are not causally implicated in the nucleation of filopodial actin
filaments. Nevertheless, these data clearly show that even though Ena/VASP proteins
contribute to filopodia formation in B16-F1 cells lacking CP, they are clearly dispensable for
the generation of these structures.

Cells 2023,12, 890 13 of 25
Cells 2023, 12, x FOR PEER REVIEW 13 of 27
3.4. CP-Deficent NIH 3T3 Cells form Numerous Filopodia on Poorly Adhesive Substrates
The strongly amplified formation of integrin-based cell adhesions, as well as the ex-
cessive spreading of NIH 3T3-derived CP-KO mutants on fibronectin, suggested en-
hanced contractility of FA-anchored SFs, likely resulting in cells with highly stretched
membranes. Interestingly, it has recently been shown that actin-based protrusions such as
lamellipodia adapt to changes in membrane tension imposed by branched actin networks,
such that large lamellipodia are formed at low tension, whereas their formation is per-
turbed at high tension [53]. Notably, we found that lamellipodia formation in NIH 3T3
fibroblasts was suppressed after loss of CP, as evidenced by immunostaining the cells
with the WRC component WAVE2 (Figure S5). Thus, in addition to the relevance of CP
for lamellipodium formation, this could also mean that filopodia formation in NIH 3T3-
derived CP-KO cells on fibronectin is suppressed because the force generated by nascent
filopodia is not sufficient to overcome the high tension of the membrane. Since it is not
possible to distinguish between these two possibilities without elaborate biophysical
methods such as atomic force microscopy (AFM) or the use of novel tension sensors
probes reporting membrane tension changes through their fluorescence lifetime [54], we
therefore asked whether filopodium formation could be induced by seeding the cells on
poorly adhesive substrates. As shown in Figure 4, CP-KO cells seeded on poly-L-lysine or
directly on glass adhered poorly but formed numerous filopodia, comparable to CP-defi-
cient B16-F1 cells plated on laminin. Thus, the striking filopodia phenotype upon loss of
CP appears to be cell-type- and context-dependent.
Figure 4.
CP-deficient NIH 3T3 fibroblasts form numerous filopodia on poorly adhesive substrates.
Representative images of NIH 3T3 wild-type and derived CP-KO mutant cells seeded on poly-L-lysine-
coated glass (
top
) or directly on glass (
bottom
) and stained for the F-actin cytoskeleton with fluorescent
phalloidin. Brightest point projection of 3D reconstructions from Airyscan confocal sections are shown.
3.6. Endogenous MyoX, FMNL2 and -3, but Not mDia2, Reside at the Tips of Filopodia in CP-KO Cells
These results clearly showed that in the absence of CP and Ena/VASP, additional
factors must contribute to filopodium formation in the mutants. According to our current
state of knowledge, besides unconventional MyoX [
34
,
55
–
57
], several formins, in particular,
mDia2 [
20
,
21
,
34
], and the formin-like family members 2 (FMNL2) and 3 [
22
,
24
,
58
,
59
] were
reported to localize at filopodia tips and have been implicated in driving these protru-
sions. We therefore asked which of these potent filopodia inducers localize at the tips
of filopodia in EVM/CP-KO mutants. To this end, we used affinity-purified polyclonal
antibodies and examined localization of these factors by indirect immunofluorescence in
EVM/CP-KO cells. Strikingly, contrary to our expectation, endogenous mDia2 was not
detected at filopodia tips in the mutant cells (Figure 6A). To exclude the possibility that
the antibody is not capable of detecting mDia2 at filopodia tips by immunofluorescence,
we ectopically expressed a constitutive variant of mDia2 lacking its autoinhibitory domain
(mDia2
∆
DAD) fused to EGFP, followed by immunolabeling with the mDia2 polyclonal
antibody. Consistent with previous work [
20
,
21
], expression of active mDia2 triggered the
formation of numerous filopodia. Most importantly, even in transfected cells expressing
very low levels of the fusion protein, as assessed by EGFP fluorescence, active mDia2 was
robustly detected at filopodia tips by the polyclonal antibody (Figure 6B), confirming its
usefulness and specificity. Since ectopically expressed active mDia2 fused to EGFP was
previously also reported to localize to the cell front and the tips of microspikes [
20
,
21
], we

Cells 2023,12, 890 14 of 25
also examined localization of endogenous mDia2 in B16-F1 cells forming prominent lamel-
lipodia. However, once again, endogenous mDia2 was neither detectable at the periphery
of the leading edge nor at microspike tips (Figure 6C). Notwithstanding this, and despite
the lack of mDia2 localization in B16-F1 cells during the interphase, endogenous mDia2
localized prominently together with tubulin at the mitotic spindle during prometaphase
and the midbody during telophase (Figure 6D). These findings were corroborated in hu-
man HeLa cells and NIH 3T3 fibroblasts (Figure S6), supporting the view that mDia2 is
rather implicated in cell division and not in the formation of filopodia. On the other hand,
endogenous MyoX as well as FMNL2 and -3 were found to accumulate markedly at the
tips of filopodia in EVM/CP-KO cells, with FMNL2 and -3 also detectable in dispersed
clusters along the shafts of filopodial actin bundles (Figure 7).
Cells 2023, 12, x FOR PEER REVIEW 14 of 27
Figure 4. CP-deficient NIH 3T3 fibroblasts form numerous filopodia on poorly adhesive substrates.
Representative images of NIH 3T3 wild-type and derived CP-KO mutant cells seeded on poly-L-
lysine-coated glass (top) or directly on glass (bottom) and stained for the F-actin cytoskeleton with
fluorescent phalloidin. Brightest point projection of 3D reconstructions from Airyscan confocal sec-
tions are shown.
3.5. Loss of CP in Ena/VASP-Deficient B16-F1 Cells Does Not Abrogate Filopodium Formation
In previous work, it has been hypothesized that Ena/VASP proteins are obligatory
for filopodium formation in the absence of CP [39]. To confirm or challenge this hypothe-
sis, we first examined VASP localization in CP-deficient B16-F1 cells. CP-KO cells stained
for the F-actin cytoskeleton and endogenous VASP indeed showed conspicuous VASP
clusters at the distal tips of filopodia (Figure 5A). Then, we disrupted the ß-subunit of CP
by CRISPR/Cas9 in Ena/VASP/Mena-deficient B16-F1 cells (EVM-KO), recently shown to
virtually lack microspikes but still be capable of forming filopodia upon transient expres-
sion of active mDia2 and unconventional myosin-X or by pharmacological inhibition of
the Arp2/3 complex [34]. Loss of CP-ß in independent clonal cell lines, denoted as
EVM/CP-KO cells, was again validated by TIDE analysis of amplified genomic target sites
and confirmed by immunoblotting (Figures 5B and S1A). Both independent EVM/CP-KO
mutant cells (clones #4 and #14) were highly similar, but, to our surprise, they still formed
numerous filopodia, similar to the CP-KO cells, as determined by confocal Airyscan im-
aging of cells stained with phalloidin migrating on laminin (Figure 5C and Movie S6).
Quantification further revealed filopodia to be about 20% shorter in EVM/CP-KO mutants
(2.2 ± 0.5 and 2.2 ± 0.5 µm) as compared to CP-KO control cells (2.8 ± 0.8 and 2.8 ± 0.7 µm)
(Figure 5D). Notably, the quantification of the number of peripheral filopodia, on the other
hand, showed no noticeable differences between these cell lines (Figure 5E), suggesting
that Ena/VASP proteins are not causally implicated in the nucleation of filopodial actin
filaments. Nevertheless, these data clearly show that even though Ena/VASP proteins con-
tribute to filopodia formation in B16-F1 cells lacking CP, they are clearly dispensable for
the generation of these structures.
Figure 5. B16-F1 mutants continue to display massive filopodia formation even after combined loss
of all Ena/VASP proteins and CP. (
A
) A CP-deficient B16-F1 cell stained for endogenous VASP and
F-actin confirms localization of VASP at the distal tips of filopodia. Inset, enlarged image of boxed
region. (
B
) Immunoblot confirming the elimination of CP-ß in two independent EVM-KO mutants
(clones #4 and #14). Loss of the ß-subunit, in turn, led, again, to the almost complete loss of the CP-
α
-
subunit. Loading control: GAPDH. (
C
) Morphologies of representative CP-KO and EVM/CP-KO
mutant cells migrating on laminin and stained for the F-actin cytoskeleton with phalloidin. Note the
continued formation of numerous filopodia in EVM/CP-KO cells. (
D
) Quantification of filopodia
length in B16-F1 wild-type and derived CP-KO and EVM/CP-KO mutant cells. Note that B16-F1
cells, as shown in Figure 1B, primarily form microspikes and barely any filopodia that protrude
beyond the periphery of the membrane. (
E
) Quantification of the number of peripheral filopodia
in CP-KO and EVM/CP-KO mutant cells. (
D
,
E
) The boxes in box plots indicate 50% (25–75%) and
whiskers (5–95%) of all measurements, with dashed black lines depicting the medians and arithmetic
means highlighted in red. (
D
) Non-parametric Kruskal–Wallis test and Dunn’s Multiple Comparison
test and (
E
) one-way ANOVA and Tukey Multiple Comparison test were used to reveal statistically
significant differences between datasets. *** p
≤
0.001; n.s.: not significant. n: number of filopodia (
D
)
or cells (E) analyzed from three independent experiments.

Cells 2023,12, 890 15 of 25
Cells 2023, 12, x FOR PEER REVIEW 16 of 27
Figure 6. Endogenous mDia2 is not detectable at the tips of filopodia and lamellipodia but accumu-
lates prominently together with tubulin at the mitotic spindle and midbody of dividing B16-F1 cells.
(A) Representative image of an EVM/CP-KO cell stained for endogenous mDia2 and F-actin. Note
the lack of mDia2 enrichment at filopodia tips. Inset, enlarged image of boxed region. Scale bar, 20
µm. (B) Ectopically expressed EGFP-mDia2∆DAD triggers filopodia formation in B61-F1 cells, with
active mDia2 localizing to the distal tips of filopodia. Remarkably, even at low expression levels,
ectopically expressed EGFP-mDia2∆DAD was effectively recognized by the anti-mDia2 antibody.
Inset, enlarged image of boxed region. (C) Heat map of endogenous mDia2 signal intensity in a
representative B16-F1 cell during interphase, forming a prominent lamellipodium. Note the lack of
mDia2 enrichment at the lamellipodium tip. (D) Representative images of B16-F1 cells stained for
endogenous mDia2, tubulin and DNA with DAPI in prometaphase (upper panel) and telophase
(lower panel).
Figure 6.
Endogenous mDia2 is not detectable at the tips of filopodia and lamellipodia but accumulates
prominently together with tubulin at the mitotic spindle and midbody of dividing B16-F1 cells.
(
A
) Representative image of an EVM/CP-KO cell stained for endogenous mDia2 and F-actin. Note the
lack of mDia2 enrichment at filopodia tips. Inset, enlarged image of boxed region. Scale bar, 20
µ
m.
(
B
) Ectopically expressed EGFP-mDia2
∆
DAD triggers filopodia formation in B61-F1 cells, with active
mDia2 localizing to the distal tips of filopodia. Remarkably, even at low expression levels, ectopically
expressed EGFP-mDia2
∆
DAD was effectively recognized by the anti-mDia2 antibody. Inset, enlarged
image of boxed region. (
C
) Heat map of endogenous mDia2 signal intensity in a representative B16-F1
cell during interphase, forming a prominent lamellipodium. Note the lack of mDia2 enrichment at the
lamellipodium tip. (
D
) Representative images of B16-F1 cells stained for endogenous mDia2, tubulin
and DNA with DAPI in prometaphase (upper panel) and telophase (lower panel).
3.7. Loss of MyoX in EVM/CP-KO Cells Leads to an Additional Reduction of Filopodia Length
To assess the contribution of MyoX in filopodium formation in the absence of CP
and Ena/VASP, we disrupted the Myo10 gene encoding MyoX in EVM/CP-KO cells us-
ing CRISPR/Cas9-mediated genome editing. Successful disruption of the gene in inde-
pendent mutants, referred to as EVM/CP/MyoX-KO, was validated by TIDE analysis and
verified by immunoblotting (Figures 8A and S1A). Both independent EVM/CP/MyoX-KO
mutants still formed numerous filopodia in a comparable fashion, as evidenced by confocal
Airyscan imaging of phalloidin-stained cells migrating on laminin, but, notably, their filopodia
appeared to be considerably shorter as compared to those formed by EVM/CP-KO cells

Cells 2023,12, 890 16 of 25
(Figure 8B). This was substantiated by quantification of filopodium length, showing filopo-
dia of EVM/CP/MyoX-KO cells to be almost 30% shorter (1.6
±
0.3 and 1.6
±
0.3
µ
m) as
compared to EVM/CP-KO control cells (2.2
±
0.5
µ
m) (Figure 8C). However, as with the elim-
ination of CP in EVM-KO cells, quantification of the number of peripheral filopodia showed
no significant differences between parental EVM/CP-KO and derived EVM/CP/MyoX-KO
cells (Figure 8D). Since filopodia formation may be linked to stabilization by adhesions formed
with the substrate [
60
], we also examined whether the EVM/CP/MyoX-KO cells exhibit
any changes in their spreading behavior on laminin as compared to the EVM/CP-KO cells.
As shown in Figure S7, additional loss of MyoX in EVM/CP-deficient cells only marginally
affected spreading of the EVM/CP/MyoX-KO mutant cells as compared to the parental cell
line, strongly suggesting that the observed reduction of filopodia length is not correlated with
diminished adhesion, but with the lack of MyoX activity driving filopodium formation.
Cells 2023, 12, x FOR PEER REVIEW 17 of 27
Figure 7. Endogenous MyoX, FMNL2 and FMNL3 accumulate at the tips of filopodia in EVM/CP-
KO cells. Representative EVM/CP-KO cells stained for MyoX and F-actin (upper panel), for FMNL2
and F-actin (middle panel) and for FMNL3 and F-actin (lower panel). Insets, enlarged images of
boxed regions.
3.7. Loss of MyoX in EVM/CP-KO Cells Leads to an Additional Reduction of Filopodia Length
To assess the contribution of MyoX in filopodium formation in the absence of CP and
Ena/VASP, we disrupted the Myo10 gene encoding MyoX in EVM/CP-KO cells using
CRISPR/Cas9-mediated genome editing. Successful disruption of the gene in independent
mutants, referred to as EVM/CP/MyoX-KO, was validated by TIDE analysis and verified
by immunoblotting (Figures 8A and S1A). Both independent EVM/CP/MyoX-KO mutants
still formed numerous filopodia in a comparable fashion, as evidenced by confocal Airy-
scan imaging of phalloidin-stained cells migrating on laminin, but, notably, their filopodia
appeared to be considerably shorter as compared to those formed by EVM/CP-KO cells
(Figure 8B). This was substantiated by quantification of filopodium length, showing filo-
podia of EVM/CP/MyoX-KO cells to be almost 30% shorter (1.6 ± 0.3 and 1.6 ± 0.3 µm) as
compared to EVM/CP-KO control cells (2.2 ± 0.5 µm) (Figure 8C). However, as with the
elimination of CP in EVM-KO cells, quantification of the number of peripheral filopodia
showed no significant differences between parental EVM/CP-KO and derived
EVM/CP/MyoX-KO cells (Figure 8D). Since filopodia formation may be linked to stabili-
zation by adhesions formed with the substrate [60], we also examined whether the
EVM/CP/MyoX-KO cells exhibit any changes in their spreading behavior on laminin as
Figure 7.
Endogenous MyoX, FMNL2 and FMNL3 accumulate at the tips of filopodia in EVM/CP-KO cells.
Representative EVM/CP-KO cells stained for MyoX and F-actin (
upper panel
), for FMNL2 and F-actin
(middle panel) and for FMNL3 and F-actin (lower panel). Insets, enlarged images of boxed regions.
3.8. Combined Inactivation of Ena/VASP, MyoX and FMNL2 and -3 Is Required to Drastically
Impair Filopodium Formation in CP-Deficient Cells
To assess the contribution of FMNL-family formins to filopodium formation, we finally
disrupted the FMNL2 and FMNL3 genes in EVM/CP/MyoX-KO cells using CRISPR/Cas9
technology. We initially isolated independent clonal cell lines with disrupted expression of
FMNL3 alone (EVM/CP/MyoX/FMNL3-KO) followed by additional disruption of FMNL2

Cells 2023,12, 890 17 of 25
(EVM/CP/MyoX/FMNL2/3-KO). In total, two independently generated double-KO clones
of each genotype, as validated by TIDE analysis and immunoblotting (Figures 9A and S1A),
were further analyzed. In contrast to the EVM/CP/MyoX-KO reference, filopodia formation
was severely impaired phenotypically in all FMNL-deficient mutants, with cells forming
only stub-like projections that barely extended beyond the cell periphery, as revealed by
imaging of phalloidin-stained cells (Figure 9B). Consistently, quantification revealed filopodia
to be 1.1
±
0.2
µ
m, more than 30% shorter in all FMNL-deficient mutants as compared
to EVM/CP/MyoX-KO control cells, at 1.6
±
0.30
µ
m (Figure 9C). Notably, analyses of
phalloidin-stained cells further showed all FMNL-KO mutant cells forming roughly 30%
less filopodia compared to control (Figure 9D). Thus, the removal of FMNL-family formins
reduced both the length and the number of filopodia, which is consistent with their ability to
nucleate and actively elongate actin filaments
in vitro
[
22
]. Unexpectedly, combined loss of
FMNL3 and FMNL2 did not cause a noticeably stronger phenotype regarding filopodium
formation than loss of FMNL3. Nevertheless, this is consistent with previously published
work analyzing protrusion and cell migration in B16-F1 cells showing that loss of FMNL2
alone generally caused weaker phenotypes as compared to loss of FMNL3 [23].
Cells 2023, 12, x FOR PEER REVIEW 18 of 27
compared to the EVM/CP-KO cells. As shown in Figure S7, additional loss of MyoX in
EVM/CP-deficient cells only marginally affected spreading of the EVM/CP/MyoX-KO
mutant cells as compared to the parental cell line, strongly suggesting that the observed
reduction of filopodia length is not correlated with diminished adhesion, but with the lack
of MyoX activity driving filopodium formation.
Figure 8. Loss of MyoX in EVM/CP-KO cells leads to additional reduction in filopodia length. (A)
Immunoblot confirming elimination of MyoX in two independent EVM/CP/MyoX-KO mutants
(clones #4-3 and #14-3). Loading control: GAPDH. (B) Morphologies of representative EVM/CP-KO
and EVM/CP/MyoX-KO cells migrating on laminin and stained for the F-actin cytoskeleton with
phalloidin. (C) Comparison of filopodium length in EVM/CP-KO versus EVM/CP/MyoX-KO cells.
Note the significantly shorter filopodia in EVM/CP/MyoX-KO mutants. (D) Quantification of the
number of peripheral filopodia in EVM/CP-KO and EVM/CP/MyoX-KO cells. (C,D) The boxes in
box plots indicate 50% (25–75%) and whiskers (5–95%) of all measurements, with dashed black lines
depicting the medians and arithmetic means highlighted in red. (C) Non-parametric Kruskal–Wallis
test and Dunn’s Multiple Comparison test and (D) one-way ANOVA and Tukey Multiple Compar-
ison test were used to reveal statistically significant differences between datasets. *** p ≤ 0.001; n.s.:
not significant. n: number of peripheral filopodia (C) or cells (D) analyzed from three independent
experiments.
3.8. Combined Inactivation of Ena/VASP, MyoX and FMNL2 and -3 Is Required to Drastically
Impair Filopodium Formation in CP-Deficient Cells
To assess the contribution of FMNL-family formins to filopodium formation, we fi-
nally disrupted the FMNL2 and FMNL3 genes in EVM/CP/MyoX-KO cells using
CRISPR/Cas9 technology. We initially isolated independent clonal cell lines with dis-
rupted expression of FMNL3 alone (EVM/CP/MyoX/FMNL3-KO) followed by additional
disruption of FMNL2 (EVM/CP/MyoX/FMNL2/3-KO). In total, two independently gener-
ated double-KO clones of each genotype, as validated by TIDE analysis and immunoblot-
ting (Figures 9A and S1A), were further analyzed. In contrast to the EVM/CP/MyoX-KO
Figure 8.
Loss of MyoX in EVM/CP-KO cells leads to additional reduction in filopodia length.
(
A
) Immunoblot confirming elimination of MyoX in two independent EVM/CP/MyoX-KO mutants
(clones #4-3 and #14-3). Loading control: GAPDH. (
B
) Morphologies of representative EVM/CP-KO
and EVM/CP/MyoX-KO cells migrating on laminin and stained for the F-actin cytoskeleton with phal-
loidin. (
C
) Comparison of filopodium length in EVM/CP-KO versus EVM/CP/MyoX-KO cells. Note
the significantly shorter filopodia in EVM/CP/MyoX-KO mutants. (
D
) Quantification of the number
of peripheral filopodia in EVM/CP-KO and EVM/CP/MyoX-KO cells. (
C
,
D
) The boxes in box plots
indicate 50% (25–75%) and whiskers (5–95%) of all measurements, with dashed black lines depicting
the medians and arithmetic means highlighted in red. (
C
) Non-parametric Kruskal–Wallis test and
Dunn’s Multiple Comparison test and (
D
) one-way ANOVA and Tukey Multiple Comparison test were
used to reveal statistically significant differences between datasets. *** p
≤
0.001; n.s.: not significant. n:
number of peripheral filopodia (C) or cells (D) analyzed from three independent experiments.

Cells 2023,12, 890 18 of 25
Cells 2023, 12, x FOR PEER REVIEW 19 of 27
reference, filopodia formation was severely impaired phenotypically in all FMNL-defi-
cient mutants, with cells forming only stub-like projections that barely extended beyond
the cell periphery, as revealed by imaging of phalloidin-stained cells (Figure 9B). Consist-
ently, quantification revealed filopodia to be 1.1 ± 0.2 µm, more than 30% shorter in all
FMNL-deficient mutants as compared to EVM/CP/MyoX-KO control cells, at 1.6 ± 0.30
µm (Figure 9C). Notably, analyses of phalloidin-stained cells further showed all FMNL-
KO mutant cells forming roughly 30% less filopodia compared to control (Figure 9D).
Thus, the removal of FMNL-family formins reduced both the length and the number of
filopodia, which is consistent with their ability to nucleate and actively elongate actin fil-
aments in vitro [22]. Unexpectedly, combined loss of FMNL3 and FMNL2 did not cause a
noticeably stronger phenotype regarding filopodium formation than loss of FMNL3. Nev-
ertheless, this is consistent with previously published work analyzing protrusion and cell
migration in B16-F1 cells showing that loss of FMNL2 alone generally caused weaker phe-
notypes as compared to loss of FMNL3 [23].
Figure 9.
Additional inactivation of FMNL-family formins in EVM/CP/MyoX-KO cells affected
not only the length but also the number of formed filopodia. (
A
) Immunoblot confirming
the elimination of FMNL3 or FMNL2 and -3 in two independent EVM/CP/MyoX/FMNL3 or
EVM/CP/MyoX/FMNL2/3 mutants (clones #4-3-4 and #14-3-5 or #4-3-4-1 and #14-3-5-2, respectively).
The asterisk indicates a nonspecific band, as shown in more detail in Figure S8. Loading control:
GAPDH. (
B
) Representative images of mutant cells indicated migrating on laminin and stained for
the F-actin cytoskeleton with phalloidin. (C) Comparison of filopodium length in respective cell line.
(
D
) Quantification of the number of peripheral filopodia in mutants as indicated. (
C
,
D
) The boxes in
box plots indicate 50% (25–75%) and whiskers (5–95%) of all measurements, with dashed black lines
depicting the medians and arithmetic means highlighted in red. (
C
) Non-parametric Kruskal–Wallis
test and Dunn’s Multiple Comparison test and (
D
) one-way ANOVA and Tukey Multiple Comparison
test were used to reveal statistically significant differences between datasets. *** p
≤
0.001; n.s.: not
significant. n: number of filopodia (C) or cells (D) analyzed from three independent experiments.

Cells 2023,12, 890 19 of 25
4. Discussion
In this work, we examined the consequences of CP loss in mouse B16-F1 melanoma
cells and NIH 3T3 fibroblasts. Consistent with previous work using RNA interference or
genetic knockout in various cell types [
39
,
48
–
50
], disruption of CP-ß expression in B16-
F1 cells was associated with markedly reduced levels of the CP-
α
-subunit and resulted
in unleashed actin assembly accompanied with drastic changes in cell morphology and
organization of the actin cytoskeleton. Notably, quantification of the total F-actin content
determined by flow cytometry of phalloidin-stained cells revealed a 4- to 5-fold higher
level in B16-F1-derived CP-KO mutants as compared to B16-F1 control. Even though
increased F-actin levels have been also detected in other cell types upon perturbation of
CP function [
49
,
50
], this value is significantly higher than the only 2-fold higher value
previously reported for CP-depleted B16-F10 cells [
49
]. This suggests that the difference is
presumably mainly due to incomplete removal of CP by siRNA interference, albeit it may
also be affected by the different quantification methods. Given the typical ratio of F-actin to
G-actin of approximately 1:1 in non-muscle cells [
50
,
61
,
62
], our data revealing a ratio of 7:1
in CP-KO cells further suggested a significant depletion of the polymerization-competent
pool of monomeric actin in the mutants. Interestingly, we additionally found increased
global actin levels in independent CP-KO cells that were almost 3-fold higher as compared
with the B16-F1 wild type, suggesting that loss of CP elicits compensatory mechanisms
to account for diminished G-actin levels by amplified expression of actin, for instance, by
increased transcription or stability of the respective mRNA. Since actin polymerization and
disassembly of actin filament plays a critical role in protrusion and cell migration [
63
,
64
], it
was not surprising to find that CP-deficient NIH 3T3 and B16-F1 mutants exhibited very
strong migration defects. These results are consistent with previous work analyzing the
motility of CP-depleted B16-F10 and Dictyostelium cells [
49
,
50
], but, again, the motility
defects were more severe upon complete loss of CP. These results are furthermore in line
with a very recent study showing that the rate of lamellipodium protrusion is considerably
impaired in CP-deficient B16-F1 cells [48].
The depletion of CP in different cell types such as B16-F1, B16-F10 and Dictyostelium
cells is commonly linked to excessive formation of filopodia [
39
,
49
,
50
], as shown here
for CP-deficient B16-F1 cells. However, in contrast to a previous study proposing that
CP-depleted NIH 3T3 fibroblasts also form numerous filopodia [
39
], we found that loss of
CP in independent clonal cell lines derived from NIH 3T3 fibroblasts was accompanied
by excessive formation of long ventral SFs linked to exaggerated FAs, occupying the
entire cell area, instead of the formation of numerous filopodia. This raises the important
question of as to why CP-deficient fibroblasts form an excess of adhesive and contractile
structures at the expense of filopodia. Of note, it has been recently shown that different
phenotypes, referred to as “clonal variability” in genome-edited cells, are already partially
attributable to the heterogeneity of wild-type cells [
65
]. Intriguingly, this study found
hundreds of differentially regulated transcripts when comparing clonal populations of
mIMCD-3 wild-type cells. Since these differences are certainly even more pronounced in
diverse cell types such as the B16-F1 melanoma cells and NIH 3T3 fibroblasts used here,
the specific phenotype that emerges after the loss of CP in one cell type cannot simply be
transferred to the other. Interestingly, ectopically expressed CP fused to GFP has been found
to localize along SFs in Xenopus XTC fibroblasts [
66
]. Since CP binds to the barbed end with
subnanomolar affinity [
67
], thus preventing the addition and loss of actin monomers at the
end, it is not surprising in our view that the loss of CP in NIH 3T3 fibroblasts promotes
amplified growth of SF filaments to generate excess of exceptionally long SFs connected to
enlarged FAs. Given the strongly amplified formation of FAs and SFs and the excessive
spreading of NIH 3T3-derived CP-KO mutants on fibronectin, we assume that enhanced
contractility of FA-anchored SFs likely leads to cells with a highly stretched membrane
that has been shown to suppress the formation of protrusive structures [
53
]. Although this
needs to be confirmed by direct measurements using AFM or membrane tether pulling,
our assumption is supported by the analysis of primary adult fibroblasts derived from

Cells 2023,12, 890 20 of 25
CP-ß-deficient mice showing that inactivation of CP leads to increased contractility and cell
tension on stiff hydrogels [
68
]. This suggests that filopodia formation in NIH 3T3-derived
CP-KO cells on fibronectin is suppressed because the pushing force of emerging filopodia is
insufficient to overcome the high tension of the membrane. Consistent with this notion, we
were able to induce the formation of numerous filopodia by seeding these CP-KO mutants
on poorly adhesive substrates.
The most important aspect of this study was the experimental reassessment of the
previous hypothesis proposing that Ena/VASP proteins are essential for filopodium forma-
tion in the absence of CP [
39
]. Even though we could confirm the explosive formation of
filopodia in CP-deficient B16-F1 mutants, we still found comparable numbers of filopodia
that were only 20% shorter in independent mutants lacking CP and all three Ena/VASP
proteins (Table 1). Thus, despite providing supportive evidence for the contribution of
Ena/VASP proteins to filopodia formation in this cell type, these results clearly challenge
the previous hypothesis by showing that additional factors must be at play that drive
the formation of these finger-shaped structures in the absence of CP and Ena/VASP. In
a systematic search for these factors, we identified endogenous MyoX and the FMNL-
family formins FMNL2 and FMNL 3 to reside prominently at the tips of distal filopodia in
EVM/CP-KO cells. Consecutive elimination of these factors resulted in a gradual further
decrease of filopodia length by 50% (Table 1), clearly demonstrating that, in addition to
Ena/VASP proteins, endogenous MyoX and FMNL3 also effectively contribute to filopodia
formation in the absence of CP. On the other hand, additional loss of FMNL2 had virtually
no effect. Given the prominent localization of endogenous FMNL2 at the tips of filopodia,
this result was rather unexpected. Nevertheless, previous work analyzing lamellipodium
dynamics and cell migration of B16-F1-derived mutant cells revealed that phenotypes
associated with the loss of FMNL2 are generally weaker than the loss of FMNL3 [
23
],
which in turn may be related to the stronger nucleation activity of FMNL3 compared to
FMNL2 [
23
]. In line with this notion, we found filopodia numbers to be decreased only in
the multiple-KO mutants lacking FMNL3 and FMNL2/3 supporting the view that filopodia
can arise independently of lamellipodial networks by formin-mediated de novo nucleation
and elongation of filopodial filaments [37,69,70].
The immunolabeling of CP-deficient B16-F1 cells further revealed that endogenous
MyoX, FMNL2 and 3 were like VASP already present at the tips of filopodia in CP-KO
cells (Figures 5A and S9), strongly supporting the view that all these factors are part of
a common machinery promoting filopodium formation in CP-deficient cells. Moreover,
cells have also frequently the remarkable ability to compensate for the loss of one protein
by up-regulating or down-regulating others. Thus, we compared global levels of MyoX
and the FMNL2/3 formins in B16-F1 cells and a set of mutants. However, we did not
detect any significant changes in the global levels of MyoX or the FMNL2/3 formins in
respective mutant cell lines compared to B16-F1 control (Figure S10). Combined, these
findings therefore not only exclude compensatory mechanisms leading to upregulation
of specific tip factors upon loss of others, but also comprehensibly explain the gradual
decrease in filopodia formation upon successive elimination of individual components.
Even though ectopic expression of constitutively-activated variants of mDia2 fused to
EGFP is well established to faithfully induce numerous filopodia-like structures, with the
formin accumulating at their tips [
21
,
34
,
71
], to our surprise, we could not detect endogenous
mDia2 at the tips of filopodia in EVM/CP-KO cells. Moreover, mDia2 could also not be
localized at other protrusive structures such as the tips of lamellipodia or microspikes in
B16-F1 cells. Given the specificity and sensitivity of the antibody, capable of detecting even
minute amounts of ectopically expressed mDia2, the diffuse staining pattern rather suggests
that mDia2 is largely autoinhibited during interphase, as release from autoinhibition has
previously been shown to be critical for subcellular localization of various Drfs [
20
,
72
–
74
].
In marked contrast, we found that endogenous mDia2 accumulates prominently along
with tubulin at the mitotic spindle and the midbody in dividing cells of different types,
supporting its critical function in cytokinesis, as already suggested by previous work [
75
–
77
].

Cells 2023,12, 890 21 of 25
These studies have shown endogenous mDia2 to be associated with the midbody and the
equatorial cortex, albeit the latter localization was not seen in our experiments. The reason
for this partly different localization pattern is currently unclear, but could be due to the
different fixation methods used. The function of mDia2 in cytokinesis is further aided by an
abnormal configuration of the spindle in mDia2-depleted cells [
78
] as well as the analysis of
mice with global knockout of mDia2, which proved to be embryonically lethal [
77
]. Thus,
although we cannot formally exclude the presence of minute amounts of mDia2 on filopodia
tips that are below the detection limit of antibody staining, collectively our data do not
support a contribution of this formin to filopodia formation.
Of note, not even the combined elimination of CP, Ena/VASP proteins, MyoX, and
FMNL-family formins was sufficient to completely abolish filopodia, raising the question
about the remaining factors promoting the generation of the short, stub-like protrusions
in the multiple KO mutants. We envision three possible scenarios to explain these results.
Firstly, residual filopodia formation could be facilitated by other, yet specific tip factors
that have not been analyzed in this study. This group could include other unconventional
barbed-end directed myosins such as Myosin IIIa, VIIa and XVa [
79
–
82
] as well as other
formins. One potential formin that has been implicated in filopodia formation is mDia1,
although conflicting results have been reported in different studies. Whereas expression of
constitutively active mDia1 for instance was found to induce filopodia-like protrusions in
NIH 3T3 cells [
74
], another study using Jurkat T lymphocytes, 300.19 pre-B lymphomas
and NIH 3T3 cells reached the opposite conclusion [
24
]. The overexpression mDia3 has
been linked to IRSp53-driven filopodium formation in N1E115 cells [
83
], even though
endogenous mDia3 could not be detected at filopodial tips. Notably, we did not detect
endogenous mDia1 and mDia3 at the distal tips of filopodia in EVM/CP-KO cells with our
specific antibodies (data not shown), suggesting that these formins do not contribute to
filopodia formation, at least in this cell type. This notion is further supported by recent work
showing that mDia1 and mDia3 are primarily involved in establishment and maintenance
of the actin-rich cell cortex in the rear [
84
]. The last member of the formin family currently
thought to contribute to filopodia formation is Daam1 [
85
,
86
]. However, in variance to
other formins that localize specifically at the tips, Daam1 is an actin-bundling protein that
localizes along filopodial shafts [
85
]. Secondly, it is also conceivable that other elongation
factors, e.g., formins not normally involved in filopodia formation, occupy free barbed ends
after the loss of specific filopodial factors to promote filament growth and thus give rise to
filopodia. Notably, filopodia can also form by convergent elongation and coalescence of
lamellipodia filaments [
87
]. Thus, as the third possibility we should also consider Arp2/3
complex-meditated actin assembly and distributive actin polymerase activity of the WRC,
shown to accelerate elongation of uncapped actin filaments [
88
], which could be followed
by bundling of the filaments by cross-linkers such as fascin, espin [
89
] or Daam1 [
85
],
leading to short and stiff bundles capable of protruding beyond the cell periphery. Thus,
the contribution all these factors in filopodium formation in the absence of CP will only be
revealed by additional combinatorial loss of function studies and phenotypic side-by-side
comparisons in future work.
Table 1.
Comparison of length and number of peripheral filopodia in analyzed mutant cell lines.
Values indicate mean
±
SD of pooled data from independent clones of respective genotype. n:
number filopodia 1or cells 2analyzed.
Mutant Cell Lines Filopodia
Length (µm)
Normalized to
CP-KO (%) n1Number of
Filopodia/Cell
Normalized to
CP-KO (%) n2
CP-KO 2.8 ±0.7 100.0 ±25.6 208 89.9 ±14.3 100.0 ±15.9 59
EVM/CP-KO 2.2 ±0.5 78.6 ±24.5 204 90.1 ±14.2 100.1 ±15.7 54
EVM/CP/MyoX-KO 1.6 ±0.3 57.7 ±20.6 206 89.0 ±11.4 99.0 ±12.8 60
EVM/CP/FMNL3-KO 1.1 ±0.2 39.7 ±20.7 198 64.2 ±11.7 71.4 ±18.3 60
EVM/CP/MyoX/FMNL2/3-KO 1.1 ±0.9 40.0 ±16.8 194 62.0 ±10.8 69.0 ±17.3 60

Cells 2023,12, 890 22 of 25
Supplementary Materials:
The following are available online at https://www.mdpi.com/article/
10.3390/cells12060890/s1, Figure S1. TIDE sequence trace decomposition analysis of clonal cell
lines derived from B16-F1 cells. Figure S2. Loss of CP in B16-F1 cells prevents the formation of
smooth lamellipodia and markedly decreases cell polarity. Figure S3. Loss of CP in B16-F1 cells
increases global actin levels. Figure S4. Loss of CP in NIH 3T3 fibroblasts increases length and
width of FAs. Figure S5. Loss of CP in NIH 3T3 fibroblasts suppresses lamellipodium formation.
Figure S6
. Endogenous mDia2 accumulates prominently together with tubulin at the mitotic spindle
and midbody of dividing NIH 3T3 and HeLa cells. Figure S7. Loss of MyoX in EVM/CP cells does
not noticeably affect cell spreading. Figure S8. Additional evidence illustrating that the cross-reacting
band shown in Figure 9A is nonspecific. Figure S9. Endogenous MyoX, FMNL2 and FMNL3 are
already present at the tips of filopodia in CP-KO cells. Figure S10. Global expression of FMNL3,
FMNL2 and MyoX in representative clonal cell lines. Figure S11. Uncropped immunoblots and
Coomassie-stained gels. Movie S1: Loss of CP in B16-F1 cells triggers the massive formation of
filopodia. Movie S2. Loss of CP in B16-F1 cells markedly perturbs lamellipodium dynamics and
induces the formation of numerous filopodia. Movie S3: Filopodia in CP-deficient B16-F1 cells are
highly dynamic. Movie S4. Loss of CP in B16-F1 cells severely impairs cell migration. Movie S5. Loss
of CP in NIH 3T3 fibroblasts also impairs cell migration. Movie S6: B16-F1 cells lacking CP and all
three Ena/VASP proteins still form prominent filopodia.
Author Contributions:
Conceptualization, J.I.H. and J.F.; methodology, J.I.H., J.S., S.K. and T.K.;
validation, J.I.H., J.S., T.K. and S.K.; formal analysis, J.I.H., J.S., T.K. and S.K.; investigation, J.I.H.,
J.S., T.K., S.K. and J.F.; resources, J.F.; data curation, J.I.H., J.S., S.K. and T.K.; software, J.S., writing—
original draft preparation, J.F.; writing—review and editing, J.F.; visualization, J.I.H., J.S. and J.F.;
supervision, J.F.; project administration, J.F.; funding acquisition, J.F. All authors have read and
agreed to the published version of the manuscript.
Funding:
This research was funded by the Deutsche Forschungsgemeinschaft, grant numbers
FA330/9-2, FA330/12-1 and FA330/13-1.
Institutional Review Board Statement:
The immunization of rabbits for the generation of polyclonal
antibodies was conducted in accordance with national guidelines for the care and maintenance of
laboratory animals and approved by the Hannover Medical School Institutional Animal Care Facility
and the Lower Saxony State Office for Consumer Protection and Food Safety (LAVES) under the
application number 18A255 to J.F.
Informed Consent Statement: Not applicable.
Data Availability Statement:
All study data are included in this article. Constructs and macros
presented in this study are available from the corresponding author upon reasonable request.
Acknowledgments: We thank Annette Breskott for excellent technical assistance.
Conflicts of Interest: The authors declare no conflict of interest.
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