Human Ribosomal G-Quadruplexes Regulate Heme Bioavailability

Abstract

The in vitro formation of stable G-quadruplexes (G4s) in human ribosomal RNA (rRNA) was recently reported. However, their formation in cells and their cellular roles were not resolved. Here, by taking a chemical biology approach that integrates results from immunofluorescence, G4 ligands, heme affinity reagents, and a genetically encoded fluorescent heme sensor, we report that human ribosomes can form G4s in vivo that regulate heme bioavailability. Immunofluorescence experiments indicate that the vast majority of extra-nuclear G4s are associated with rRNA. Moreover, titrating human cells with a G4 ligand alters the ability of ribosomes to bind heme and disrupts cellular heme bioavailability as measured by a genetically encoded fluorescent heme sensor. Overall, these results suggest ribosomes play a role in regulating heme homeostasis.

Full text

Human ribosomal G-quadruplexes regulate heme
bioavailability
Receivedfor publication,May 12, 2020, and in revised form, August 6, 2020 Published, Papers in Press, August13, 2020, DOI 10.1074/jbc.RA120.014332
Santi Mestre-Fos
1,2,3
,Chieri Ito
1,2,3
,Courtney M. Moore
2,3
,Amit R. Reddi
2,3,4,
*, and Loren Dean Williams
1,2,3,4,
*
From the
1
Center for the Origin of Life,
2
School of Chemistry and Biochemistry,
3
Parker Petit Institute of Bioengineering and
Biosciences, and
4
School of Biological Sciences, Georgia Institute of Technology, Atlanta, Georgia, USA
Edited by Karin Musier-Forsyth
The in vitro formation of stable G-quadruplexes (G4s) in
human rRNA was recently reported. However, their formation
in cells and their cellular roles were not resolved. Here, by tak-
ing a chemical biology approach that integrates results from im-
munofluorescence, G4 ligands, heme-affinity reagents, and a
genetically encoded fluorescent heme sensor, we report that
human ribosomes can form G4s in vivo that regulate heme bioa-
vailability. Immunofluorescence experiments indicate that the
vast majority of extra-nuclear G4s are associated with rRNA.
Moreover, titrating human cells with a G4 ligand alters the abil-
ity of ribosomes to bind heme and disrupts cellular heme bio-
availability as measured by a genetically encoded fluorescent
heme sensor. Overall, these results suggest that ribosomes play
a role in regulating heme homeostasis.
Heme (iron protoporphyrin IX) is an essential butpotentially
cytotoxic metallocofactor and signaling molecule required for
much of life on Earth. All heme-requiring cells and organisms
must tightly regulate heme concentration and bioavailability
to mitigate toxicity (1–4). Proteins that synthesize and
degrade heme are relatively well-understood; structures and
mechanisms of all eight heme biosynthetic enzymes and heme-
degrading heme oxygenases are known (2–4). However, regula-
tion of heme bioavailability, including its intracellular traffick-
ing from sites of synthesis in the mitochondrial matrix or
uptake at the plasma membrane, is poorly understood. Current
paradigms for heme trafficking and mobilization involve heme
transfer by unknown proteinaceous factors and largely ignore
contributions from nucleic acids. Given that the first opportu-
nity for protein hemylation occurs during or just after transla-
tion, rRNA or ribosomal proteins (rProteins) may be critical for
shepherding labile heme to newly synthesized proteins.
We hypothesize that heme bioavailability is regulated in part
by ribosomes via rRNA G-tracts, which are continuous runs of
guanines. G-tracts are confined primarily to ribosomes of birds
and mammals (5) and are focused in rRNA tentacles, which are
seen to extend for hundreds of Å from ribosomal surfaces in
these species (6). Tentacles are elaborations of rRNA expansion
segments, which help form the secondary shell around the
common core of eukaryotic ribosomes (7).
Tandem G-tracts can form G-quadruplexes (G4s), which are
nucleic acid secondary structures composed of four guanine
columns surrounding a central cavity that sequesters mono-
valent cations. Our rRNA
G4
-heme hypothesis is based in
part on our observation of stable rRNA G4s in vitro (5,8)
and the extraordinary abundance of rRNA in vivo (9). Our
rRNA
G4
-hemehypothesisisalsobasedonworkbySenand
co-workers (10–12), who has demonstrated high affinity of
heme for G4s (K
D
;nM)andproposedthatRNAandDNA
G4s sequester heme in vivo (13).
DNA G4s are thought to help regulate replication (14), tran-
scription (15), and genomic stability (16). In mRNA, G4s are
associated with untranslated regions and have been proposed
to regulate translation (17–19). However, the in vivo folding
state and functional roles of G4s are under debate. It has been
proposed that eukaryotic RNA G4s are unfolded by helicases
(20), although some investigators are not convinced (21,22).
The density of G4 sequences on surfaces of the human ribo-
some, which is extremely abundant, is high, with 17 G4 sequen-
ces in the 28S rRNA and three in 18S rRNA (Fig. 1A). Previous
to this report, it was not known whether human ribosomes
form G4s in vivo or what their functions might be.
Here we present evidence that human rRNA tentacles form
G4s in vivo that regulate cellular heme homeostasis. Results of
immunofluorescence experiments with a G4 antibody, RNA
pulldowns, and experiments with well-characterized G4 ligands
provide strong support for in vivo formation of surface-exposed
G4s on rRNA tentacles. We find that G4s on ribosomes bind
heme in vitro (Fig. 1B) and that perturbation of G4s in vivo with
G4 ligands affects heme interactions and bioavailability, as meas-
ured by heme-affinity reagents and genetically encoded heme
sensors. The effects of in vivo G4-heme perturbations are pre-
dicted by in vitro experiments. Taken together, the results here
indicate that rRNA G4s interact with heme in cells and suggest
that ribosomal G4s play roles in intracellular heme metabolism.
Results
rRNA forms G4s in vivo
Confocal microscopy and G4-pulldowns were used to deter-
mine whether human ribosomes form G4s in vivo. For confocal
microscopy, we used the BG4 antibody, which selectively targets
G4s (23,24) and has been broadly used for visualizing DNA G4s
and non-rRNA G4s in cells (24–27). Our method of permeabiliz-
ing cells for antibody treatment does not permeabilize the nuclei
(28). Therefore, DNA G4s were not anticipated or observed. To
This article contains supporting information.
*For correspondence: Loren Dean Williams, loren.williams@chemistry.
gatech.edu;AmitR.Reddi,amit.reddi@chemistry.gatech.edu.
J. Biol. Chem. (2020) 295(44) 14855–14865 14855
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identify ribosome-associated G4s, we determined the extent to
which antibodies to rProtein L19 (eL19) and to G4s colocalize
and how colocalization is altered when cells are subjected to
RNase or G4 ligand PhenDC3. Prior to antibody addition, the
cells were cross-linked with paraformaldehyde, which has been
shown to lock G4s in situ and reduce induction of G4s by small
ligands (22). The extent of L19 and G4 antibody colocalization
suggests that a fraction of ribosomes form G4s (Fig. 2, Aand C)
and that most G4s are associated with ribosomes. Specifically, we
find that ;83% of BG4 colocalizes with L19, indicating that the
vast majority of RNA G4s in vivo are associated with ribosomes
(Fig. 2C,green bar) and are therefore rRNA G4s. Conversely, only
5% of L19 colocalizes with BG4 (Fig. 2C,WT red bar), indicating
that only a specialized fraction of ribosomes contain G4s. Similar
results were obtained using an antibody against rProtein uL4
instead of eL19 (not shown). It is possible that the polymorphic
nature of rRNA G4s attenuates binding to BG4, contributing to
low colocalization ratios.
PhenDC3 (3,39-[1,10–phenanthroline-2,9-diylbis(carbonyli-
mino)]bis[1-methyl quinolinium] 1,1,1-trifluoromethanesulfo-
nate) is a bisquinolinium phenanthroline derivative known to
induce and stabilize G4s (29–32). Here, PhenDC3 appears to
increase ribosomal G4 formation in vivo;treatingcellswith
PhenDC3 increases L19-BG4 colocalization from 5 to ;24% (Fig.
2C).TheincreaseincolocalizationuponPhenDC3treatment
supports formation of G4s by ribosomes. By contrast, treating
cells with RNase A abolishes the L19-BG4 colocalization signal
(Fig. 2C). Together, these results indicate that the colocalized
BG4 signal is from a G4-forming RNA in close proximity to L19.
The high density of ribosomes on the surface of the endo-
plasmic reticulum (ER) and the lower abundance of mRNA in
this location as compared with the cytosol (33) motivated us to
investigate whether G4s colocalize with the ER. mRNAs in the
cytosol, in the unlikely event that they form G4s at high fre-
quency (20), may confound our ability to selectively detect
rRNA G4s. Toward this end, we determined the extent to
which BG4 colocalizes with an antibody against an ER mem-
brane protein (calnexin) (Fig. 2B). Indeed, we find that ;45% of
the BG4 signal colocalizes with the ER surface (Fig. 2D,green
bar), indicating a significant presence of RNA G4s near the ER
membrane. As with L19, the fraction of the ER signal that
colocalizes with G4s is completely abolished by RNase A and
enhanced by PhenDC3 (increasing from 2 to 12%) (Fig. 2D).
Thus, the data are consistent with formation of rRNA G4s by
ER-bound ribosomes.
In an orthogonal approach, we pulled down RNA with Bio-
TASQ (22,34), a G4 ligand linked to biotin that captures G4s.
We previously used BioTASQ to demonstrate that human
rRNA forms G4s in vitro (Fig. 2E)(8). Here, we captured rRNA
G4s from cross-linked HEK293 cells by methods summarized
in Fig. 2F. BioTASQ captures 28S rRNA from cell lysates (Fig.
2G), in agreement with our previous in vitro BioTASQ data and
with observations of G4-L19 colocalization above. BioTASQ
also captures 18S rRNA, although the signal is significantly
weaker. This observation is in agreement with the greater abun-
dance of G-tracts in human 28S rRNA (17 G4 regions) than in
18S rRNA (three G4 regions). Taken together, our immunoflu-
orescence and BioTASQ experiments provide strong evidence
that human ribosomes form G4s in vivo.
Human ribosomes bind hemin in vitro
It has been suggested that G4s might associate with heme in
vivo (10,11,35). In vitro, heme binds with high affinity to G4s
by end-stacking (10–12,36–38)(Fig. 1B). Here, we used UV-
visible spectroscopy to assay the binding of hemin to human
rRNA. rRNA oligomers GQES7-a (Fig. 3A), GQES7-b (Fig.
S1A), or GQes3 (Fig. S1B) were titrated into fixed amount of
hemin. GQES7-a and GQES7-b are fragments of expansion
segment 7 of human large subunit (LSU) rRNA (5). GQes3 is a
fragment of expansion segment 3 of human small subunit
(SSU) rRNA (8). Each of these oligonucleotides is known to
form G4s, and each caused a pronounced increase in the Soret
band of hemin at 400 nm. The binding is specific for G4s
because a mutant oligonucleotide, mutes3, that lacks G-tracts
does not induce a change in the hemin Soret band (Fig. S1C).
Larger human ribosomal components also bind heme. Intact
28S and 18S rRNAs extracted from human cells (Fig. S1, Dand
E), assembled LSUs (Fig. 3B) and SSUs (Fig. S1F), and poly-
somes (Fig. 3C) all induce changes in the hemin Soret bands,
which is indicative of heme-rRNA interactions. Fitting of the
UV-visible data in Fig. 3A, using a one-site binding model
yielded an apparent limiting K
D
value of ,100 nM(Fig. 3H),
which is similar to that of other G4s (10–12,39). The combined
data are consistent with a model in which rRNA tentacles of
human ribosomes bind to hemin in vitro.
PhenDC3 was used to confirm binding of hemin to ribo-
somal G4s under initial conditions that favor G4 formation.
PhenDC3, like hemin, end-stacks on G4s (30,35). In vitro,
under conditions favoring G4s (50 mMK
1
), essentially all
rRNA G-tracts form stable G4s prior to PhenDC3 addition (5,
8). Under these conditions, PhenDC3 competed with heme for
binding to rRNA G4s (Fig. 3, D–F). With fixed concentrations
of GQES7-a and hemin, additionof PhenDC3 decreased the in-
tensity of the hemin Soret peak(Fig. 3D) because of dissociation
Figure 1. A, secondary structures of the human LSU rRNAs (5.8S and 28S)
and SSU rRNA (18S). G4 sequences are highlighted in green. rRNA-based
oligomers from the LSU (GQES7-a and GQES7-b) and from the SSU (GQes3)
are indicated. B, schematic representation of a heme-G4 complex.
Human ribosomes appropriate heme
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Figure 2. rRNA G 4s in HEK293 cells. Colocalization of (A) ribosomal protein L19 or (B) endoplasmic reticulum (red)withRNAG4s(green). Nuclei were
stained with 49,6-diamidino-2-phenylindole (blue). C, extent of colocalization is quantitated as the ratio of colocalized pixels over total L19 pixels (red
bars) or as the ratio of colocalized pixels over total BG4 pixels (green bar). The same analysis was performed for ER-BG4 colocalization (D). The statistical
significance relative to WT is indicated by asterisks using an ordinary one-way analysis of variance with Dunnett’s post-hoc test. Each dot represents a
biological replicate. Images of cells treated without primary antibodies or with RNase A or PhenDC3 are shown in Fig. S6 and Fig. S7.E,theG4ligand
BioTASQ binds to 28S and 18S rRNAs in vitro. In the presence of BioTASQ and streptavidin beads, human rRNAs do not enter the native agarose gel. F,
schematic representation of the BioTASQ pulldown protocol. G, RT-qPCR analysis of rRNAs pulled down by BioTASQ. The statistical significance rela-
tive to a fold-enrichment value of 1 is indicated by asterisks using a one-sample ttest and a Wilcoxon test. Each dot represents a biological replicate.
Data in (G) are represented as RNA enrichment under BioTASQ 1streptavidin beads conditions relative to control streptavidin beads. *p,0.05. n.s.,
not significant.
Human ribosomes appropriate heme
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of heme. The same phenomenon was observed with assembled
ribosomal particles (LSU: Fig. 3E,SSU:Fig. S2A) and with poly-
somes (Fig. 3F). Hemin associated with purified 28S and 18S
rRNAs is dissociated by addition of PhenDC3 (Fig. S2, Band
C). Solutions of hemin with control RNA mutes3 do not show
a change in the Soret peak intensity upon the addition of
PhenDC3 (Fig. S2D). PhenDC3 absorbs at 350 nm (Fig. S2E),
causing a shoulder on the heme Soret band (Fig. 3, D–F). The
results here provide strong support for association of heme
with G4s of human ribosomes in vitro.
Unlike the in vitro experiments with K
1
, it seems probable
that most rRNA G-tractsare unfolded in cells. This inference is
based on our observation that only 5% of ribosomes bind to the
BG4 antibody in vivo until the addition of PhenDC3, upon
Figure 3. Human rRNA G4s bind to heme in vitro.UV-visible spectra (the heme Soret band) during heme titration under initial conditions that favor G4 for-
mation with (A) GQES7-a, (B) the assembled LSU, or (C) polysomes. UV-visible spectra during titration with PhenDC3 of (D) constant [heme] and [GQES7-a], (E)
constant [heme] and [LSU], and (F) constant [heme] and [polysomes]. G, UV-visible spectra during titration of constant [heme] and [GQES7-a] under initial con-
ditions that favor G4 unfolding. The absorbance at
l
max
versus [PhenDC3] is plotted on the panel on the right. H, plot of absorbance at 400 nm versus [GQES7-
a]. Data in (A) were fit to a one-site binding model (black line) giving an apparent limiting K
D
of ,100 nM. The experiments in panels D–Fwere performed using
initial conditions that favored G4 formation by G-tracts (50 mMK
1
with titration of PhenDC3 in the mMrange). The experiment in panel G was performed using
initial conditions that favored unfolded G-tracts (10 mMLi
1
,0K
1
with titration of PhenDC3 nMrange).
Human ribosomes appropriate heme
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which BG4 binding increases to 24% of ribosomes (Fig. 2, Cand
D). This inefficient folding of G-tracts into G4s in our in vivo
experiments is in agreement with previous studies (20).
We mimicked the in vivo environment using initial condi-
tions that favor unfolded G4s (Li
1
, low PhenDC3) by GQES7-a
rRNA. Under these conditions, we observed that PhenDC3
caused an increase in the binding of heme to rRNA G4s at con-
centrations below 25 nMPhenDC3, as inferred from an increase
in the absorbance of the heme Soret peak (Fig. 3G). However, at
PhenDC3 concentrations above 25 nM, we observed a decrease
in the binding of heme to rRNA G4s, as indicated by a reduc-
tion in the absorbance of the heme Soret band (Fig. 3G). Thus,
under initial conditions that favor unfolded G-tracts (Li
1
), low
PhenDC3 enhanced heme-binding to GQES7-a. The coopera-
tive relationship between PhenDC3 and heme under some con-
ditions is expected because multiple ligand binding sites are
formed by a single binding event (30). On the other hand, under
initial conditions that favor folding of G-tracts to G4s (K
1
),
PhenDC3 acted as a competitor of heme-binding to GQES7-a.
It seems possible that formation of G4s by the extended arrays
of G-tracts in rRNAs might be cooperative, although to our
knowledge this has not been demonstrated.
Human ribosomes bind heme in vivo
We developed an assay that exploits differential interactions
with hemin-agarose, an agarose resin covalently linked to
heme, to report in vivo heme-binding to ribosomes and rRNA.
The degree to which any biomolecule interacts with heme in
cells is inversely correlated with the extent to which it interacts
with hemin-agarose upon lysis because of competition between
endogenous heme and hemin-agarose. Therefore, the effects of
heme-binding factors in vivo can be monitored by determining
whether their interaction with hemin-agarose changes upon
depletion of intracellular heme.
Accordingly, HEK293 cells were grown with or without
succinylacetone (SA) (40), an inhibitor of heme biosynthe-
sis. Lysates of these cells were incubated with hemin-aga-
rose, and hemin-agarose–interacting rRNA was quantified
by RT-qPCR. Consistent with previous work (41), treatment
with 0.5 mMSA for 24 h caused a 7-fold decrease in total cel-
lular heme in HEK293 cells (results not shown). The results
reveal that rRNA binding to hemin-agarose relative to control
agarose lacking heme increases by ;4-fold in cells depleted of
heme (Fig. 4A). This result suggests that under heme-depleted
conditions, a greater fraction of rRNA-heme–binding sites are
free and available to bind hemin-agarose. In short, the data are
consistent with a model in which ribosomal RNAs associate
with endogenous heme.
PhenDC3 treatment of cells increases binding of ribosomes to
hemin-agarose
To probe rRNA
G4
-heme–binding in vivo, we determined
whether rRNA from HEK293 cells treated with the G4 ligand
PhenDC3 (48 h at 37 °C) would bind more extensively to he-
min-agarose. RT-qPCR reveals that PhenDC3 treatment of
HEK293 cells causes a dose-dependent increase in binding of
the LSU to hemin-agarose (Fig. 4B). A corresponding but
weaker signal is seen for the SSU, in agreement with the higher
abundance of G4 regions in the LSU than in the SSU (Fig. 1A).
These data are consistent with our observations that PhenDC3
promotes rRNA G4 formation in cells (Fig. 2, C–D), providing
additional heme-binding sites that can interact with hemin-
agarose. Control experiments show that PhenDC3 as used here
does not alter rRNA levels (Fig. S4) and that carrier DMSO
does not affect the results (Fig. S3, Band C).
rRNA G4s regulate heme bioavailability in vivo
To determine whether rRNA G4s regulate heme homeosta-
sis, we deployed a previously described genetically encoded
ratiometric fluorescent heme sensor, HS1. HS1 is a tri-domain
fusion protein consisting of heme-binding domain cytochrome
b
562
fused to two fluorescent proteins. The fluorescence of
eGFP is quenched by heme, and the fluorescence of mKate2 is
unaltered by heme. Thus, the ratio of eGFP:mKate2 fluores-
cence is inversely correlated with bioavailable heme, as mea-
sured by HS1. HS1 was previously used to characterize heme
homeostasis in yeast, bacteria, and mammalian cells and was
instrumental in identifying new heme-trafficking factors and
signals that alter heme biodistribution and dynamics (40,42–
44). We asked if cytosolic heme bioavailability is altered in
Figure 4. Ribosomes appropriate heme in vivo through rRNA G4s. A, RT-
qPCR analysis from untreated (WT) and SA-treated human cells. Statistical
significance relative to WT is represented by asterisks using Student’sttest.
Each dot represents a biological replicate. B, RT-qPCR analysis from PhenDC3-
treated HEK293 cells. Statistical significance relative to no-treatment condi-
tions is represented by asterisks using ordinary one-way analysis of variance
with Dunnett’s post-hoc test. Each dot represents a technical replicate com-
ing from individual biological replicates. The experiment was performed two
times with similar dose-dependent trends (Fig. S3A). Data in (A) and (B)are
represented as RNA enrichment in hemin-agarose beads relative to control
Sepharose beads. C, single-cell analysis of HS1-transfected HEK293 cells
grown in HD1SA, regular media containing 5-aminolevulinic acid (R1ALA),
or regular media (regular) with the indicated concentrations of PhenDC3. Sta-
tistical significance relative to regular conditions is represented by asterisks
using the Kruskal-Wallis analysis of variance with Dunn’s post-hoc test. *p,
0.05, **p,0.01, ***p,0.001, ****p,0.0001; n.s., not significant; n%1500
cells. D, median HS1 sensor ratios obtained in (C) as a function to PhenDC3
concentration.
Human ribosomes appropriate heme
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response to G4 ligand PhenDC3 (40). As indicated in Fig. 4C,
single-cell analysis of a population of ;1500 HEK293 cells/con-
dition indicated that the median HS1 eGFP/mKate2 ratio
increased upon heme depletion in heme-deficient media con-
taining SA (HD1SA) and decreased upon increasing intracel-
lular heme when cells were conditioned with the heme biosyn-
thetic precursor 5-aminolevulinic acid (ALA) to drive heme
synthesis. Upon exposure to PhenDC3 for 24 h, which is the
minimal amount of time needed to observe an effect on labile
heme (Fig. S5), the HS1 sensor ratio increased in response to
increasing PhenDC3 dose, indicating a decrease in heme bio-
availability. This 24-h treatment is similar to treatment times
used previously for small-molecule induction of G4s in cells
(23,24,35). The difference in the time required for PhenDC3
to affect heme in vivo (24 h) versus in vitro (15 min) is likely due
to the kinetics of PhenDC3 cellular import, its partitioning into
rRNA, and its induction of G4 formation, followed by the con-
sequent reequilibration of cellular heme into G4 heme-binding
sites. Regardless, these observations are in agreement with our
data in Fig. 3G. PhenDC3 induced G4 formation in vivo,
increasing the heme-binding sites on the rRNA that can bind
heme. Consequently, there was a decrease in the bioavailable
heme as detected by HS1. The fractional heme saturation of
HS1 decreased by ;15% (Fig. 4D), which is on the order of
what one would expect based on an equilibrium competition
model between HS1 and intracellular rRNA G4s that takes into
account the relative abundance of these species and their Fe(III)-
heme affinities (Fig. S8). Our data indicate that rRNA G4s bind
heme and regulate intracellular heme bioavailability.
Discussion
Over the last two decades, whereas a handful of proteins
have been implicated in regulating heme homeostasis on the
basis of binding heme in vitro, there has been very little evi-
dence of such roles from in-cell or in vivo studies (2–4,45).
Two notable exceptions include GAPDH and progesterone re-
ceptor membrane component 1 and 2. GAPDH, a key glycolytic
enzyme, was found to bind and buffer heme, regulatingits bioa-
vailability (40,43), and to deliver heme to nuclear heme-de-
pendent transcription factors (40) and heme enzymes such as
nitric oxide synthase (43,46) and guanylate cyclase (47). Proges-
terone receptor membrane components, which interact with the
terminal heme synthetic enzyme, ferrochelatase (48), have long
been known to bind heme and affect the activity of P450
enzymes, raising the specter that they are heme chaperones (1,
49–53). More recently, they were found to enable the delivery of
heme to the nucleus to control metabolism in adipocytes (54).
However, current paradigms for heme trafficking and mobiliza-
tion are heavily protein-centric and often ignore contributions
from nucleic acids, which are highly abundant in cells.
The results here provide strong evidence that tentacles of
human ribosomes form G4s in vivo and that these G4s are
involved in appropriating heme. Immunofluorescence experi-
ments with BG4 and L19 antibodies suggest that a specialized
fraction of cytosolic ribosomes (;5%) form G4s and that most
extra-nuclear G4s (;83%) are ribosomal. The small fraction of
ribosomes observed to form G4s in vivo contrasts with the high
stability of ribosomal G4s in vitro (5,8). This difference sup-
ports Guo and Bartel (20), who suggest that eukaryotic cells
have machinery that tends to unfold G4s. Our data show that
only a small fraction of potential G4s form in vivo and that the
fraction can be increased by G4-stabilizing ligands (Fig. 2). The
high concentration of rRNA acts in opposition to the low fre-
quency per ribosome so the RNA G4s are abundant. Moreover,
the RNA G-quadruplexome appears to be ribosome-centric.
We previously reported that surfaces of both the SSU and
the LSU contain G4 sequences (5,8). A broad variety of data
are consistent with more extensive formation of G4s on the
LSU than on the SSU. These data include
more abundant and more expansive G-tracts on the LSU
than the SSU (8),
greater conservation over phylogeny of LSU G-tracts than
SSU G-tracts (5,8),
higher thermodynamic stability of LSU G4s than SSU G4s
(5,8),
greater heme-binding to G4 oligomers from the LSU than
those from the SSU (Fig. 3Aand Fig. S1B),
greater enrichment of LSU than SSU particles in BioTASQ
pulldowns (Fig. 2G),
greater enrichment of LSU than SSU particles in hemin-
agarose pulldowns (Fig. 4A), and
greater effect of in vivo PhenDC3 treatment on LSU than on
SSU rRNA in hemin-agarose pulldowns (Fig. 4B).
Our finding that rRNA G4s associate with heme in vivo has
major implications for the physiology of G4-heme interactions.
Decades of in vitro biophysical and chemical characterizations
indicate that G4 and heme interact with high affinity (K
D
;nM)
and are potent redox catalysts, facilitating peroxidase and per-
oxygenase reactions (10–12). However, it remained unclear if
heme-G4 complexes form in vivo and if heme-G4 catalyzed
reactions were physiologically relevant.
The results of three orthogonal approaches from three
groups appear to be self-consistent. Maizels and co-workers
(35) recently proposedthat heme binds to G4s in vivo, based on
the transcriptional response of cells to PhenDC3. PhenDC3
causes up-regulation of heme-degrading enzymes such as heme
oxygenase and other iron and heme homeostatic factors. These
responses were interpreted to support a model in which G4s
sequester and detoxifyheme in cells. Work by Sen and co-work-
ers (13), concurrent with ours, established G4-heme interac-
tions in vivo by exploiting the peroxidase activity of G4-heme
complexes to self-biotinylate G4s in RNA and DNA usinga phe-
nolic-biotin derivative. Here, we demonstrate in vivo heme-
rRNA
G4
interactions by several methods. The K
D
values for
heme-rRNA
G4
complexes (;nM) are on the order of concentra-
tion of labile heme (25–300 nM)(40,55,56). Therefore, rRNA
appears to be poised to buffer labile heme. We propose that
heme-rRNA
G4
interactions may be important for protein hemy-
lation reactions and/or for buffering cytosolic heme, mitigating
its potential toxicity. It remains to be determined how endoge-
nous factors and processes may modulate G4 formation to regu-
late heme availability and homeostatic mechanisms.
Human ribosomes appropriate heme
14860 J. Biol. Chem. (2020) 295(44) 14855–14865
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Summary
The results here, building on previous in vitro work, provide
the first demonstration of in vivo formation of G4s by rRNA.
The levels of rRNA G4 formation observed in vivo support a
model in which most G-tracts are unfolded in eukaryotes (20).
Even so, the extreme density of ribosomes in vivo and the large
number of G-tracts/ribosome implies a large absolute number
of rRNA G4s in vivo. We provide evidence that rRNA G4s are
inducible by small molecules and can bind heme in vivo. The
ribosome may help regulate heme bioavailability in cells and
may be directly involved in protein hemylation. These results
provide new insights into the molecules and mechanisms
underlying intracellular heme trafficking and bioavailability,
which are currently poorly understood (1–4). Our results sug-
gest that ribosomes, G4-containing rRNAs in particular, may
regulate heme metabolism, acting to buffer intracellular heme
and possibly regulate heme trafficking and cotranslational
hemylation. The ribosome as a potential heme buffer is consist-
ent with its role as a general and versatile sink for ions and small
molecules, including antibiotics (57), platinum-based drugs
(58–60), metabolites (61), and metal cations Mg
21
,Ca
21
,
Mn
21
,Fe
21
,andK
1
(62–67).
Experimental procedures
Cell culture
HEK293 cells were cultured in DMEM containing 4.5 g/liter
glucose without sodium pyruvate and L-glutamine (Corning)
supplemented with 10% FBS (Corning) and 2% penicillin-strep-
tomycin solution (Gibco) in a humidified incubator kept at
37 °C with a 5% carbon dioxide atmosphere.
RNAs
GQES7-a and GQES7-b were synthesized in vitro by tran-
scription (HiScribe
TM
T7 High Yield RNA Synthesis Kit, New
England Biolabs). GQes3 and mutes3 were purchased from
Integrated DNA Technologies. Human 28S and 18S rRNAs
were extracted from HEK293 cells with TRIzol (Invitrogen).
Intact rRNAs were isolated by pipetting from a native agarose
gel after running the rRNA into wells in the center of the gel.
The rRNA was then precipitated with 5 Mammonium acetate-
acetic acid (pH 7.5) with excess ethanol. RNA sequences are
listed in Table S1.
RNA annealing
RNAs were annealed by heating at 95 °C for 5 min and cooled
to 25 °C at 1 °C/min and incubated for 10 min at 4 °C.
UV-visible absorbance heme-RNA binding
Stock solutions of heme chloride (1 mM) were prepared in
DMSO. Prior to use, the heme chloride solution was sonicated
for 10 min. RNAs (GQES7-a, GQES7-b, GQes3, and mutes3)
were annealed as described above in 50 mMKCl and 10 mM
Tris-HCl, pH 7.5 in increasing RNA concentrations (for rRNA
oligomers: from 0.3 to 1 equivalent of heme). The annealing
buffer for intact 28S and 18S rRNAs and assembled ribosomal
subunits and polysomes was the same as that of the rRNA
oligomers except for the inclusion of 10 mMMgCl
2
. After RNA
annealing, heme was added to a final concentration of 3 mM.
Samples (20 ml) were allowed to stand at room temperature for
30 min, then loaded onto a Corning 384-well flat clear bottom
microplate. Absorbances were recorded from 300 to 700 nm on
a BioTek Synergy
TM
H4 Hybrid plate reader.
UV-visible absorbance, heme-PhenDC3 competition/
cooperation assay
For heme-PhenDC3 competition assays, RNAs were annealed
and allowed to bind to heme as above. Final heme concentration
was 3 mM. Final RNA concentrations were GQES7-a (3 mM), intact
human 18S rRNA (65 nM), and intact human 28S rRNA (22 nM).
After solutions were incubated for 30 min at room temperature,
PhenDC3 or carrier DMSO was added to final concentrations
consisting of 1.5 mM,3mM,and6mM.Samples(20ml) were allowed
to stand at room temperature for 15 min and were loaded onto a
Corning 384-well flat clear bottom microplate. Absorbance was
recorded from 300 to 700 nm. For PhenDC3-heme cooperation
assay (data on Fig. 3G), GQES7-a RNA (1 mM) was added to 3 mM
heme in 10 mMLiCl and 10 mMTris-HCl, pH 7.5 with no RNA-
annealing step. After RNA-heme solutions were incubated for 30
min at room temperature, PhenDC3 was added to a final concen-
tration range consisting of 1.33–133 nMand allowed to mix for
15 min. The remainder of the experiment was performed as the
competition assay described above in this section.
Heme-rRNA dissociation constants were determined from
the one-site binding model (68) depicted in the below equa-
tions using nonlinear least squares regression analysis software
KaleidaGraph 4.5 (Synergy Software, Reading, PA).
rRNA 2Hm
½
5:53ðKD1rRNAT1HmT
22KD2rRNAT2HmT
ðÞ
224rRNATHmT

:5
Abs ¼Abso1DAbs 3rRNA Hm
½
=rRNA
½
T
Where K
D
is the rRNA-heme dissociation constant, Hm
T
is
the concentration of heme, rRNA-Hm is the concentration of
the rRNA-heme complex, rRNA
T
is the concentration of rRNA
that is being titrated, Abs is the absorbance at any given con-
centration of rRNA
T
, Abs
o
is the initial absorbance of heme in
the absence of rRNA, and DAbs is the change in fluorescence
due to the formation of the rRNA-heme complex.
For data fitting, Abs
o
,DAbs, rRNA
T
, and Hm
T
were treated
as fixed parameters derived from experiments, and the K
D
was
a“floating”parameter that was derived from regression analy-
sis. The absorbance signals utilized to determine K
D
were from
the heme Soret band at 400 nm.
Total heme quantification of untreated and SA-treated
HEK293 cells
Heme was quantified as described (69). Briefly, HEK293 cells
were seeded in complete DMEM media at an initial confluency
of 10% and incubated at 37 °C for 48 h. Media for SA-treated
cells was replaced by DMEM supplemented with 10% heme-
depleted FBS and 0.5 mMSA. Heme depletion of serum was
Human ribosomes appropriate heme
J. Biol. Chem. (2020) 295(44) 14855–14865 14861
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performed as described (30). Media for untreated cells was
replaced by complete media (supplemented with 10% regular
FBS) and allowed to seed at 37°C for 24 h. The cells were har-
vested by scrapping and counted using an automated TC10 cell
counter (Bio-Rad). Then, 2.5 310
4
cells/condition were treated
with 20 mMoxalic acid and incubated at 4 °C overnight in the
dark. An equal volume of 2 Moxalic acid was added to the cell
suspensions. The samples were split, with half incubated at
95 °C for 30 min and half incubated at room temperature for 30
min. The samples were centrifuged at 21,000 3gfor 2 min, 200
ml of each was transferred to a black Greiner Bio-One flat bot-
tom fluorescence plate, and porphyrin fluorescence (excitation
(ex): 400 nm, emission (em): 620 nm) was recorded on a Syn-
ergy Mx multi-modal plate reader. Heme concentration was
calculated from a standard curve prepared by diluting a 0.1 mM
hemin chloride stock solution in DMSO and treated in the
same way as the cell suspensions above. To calculate heme con-
centration, the fluorescence of the unboiled samples was taken
as the background level of protoporphyrin IX and subtracted
from the fluorescence of the boiled sample, which is used as the
free base porphyrin produced upon the release of the heme
iron. Using this method, our data suggest SA-treatment of
HEK293 cells results in a 7-fold decrease in the total cellular
heme concentration.
Hemin-agarose binding
HEK293 cells were seeded onto a 6-well plate at an initial
confluency of 20% in DMEM with 10% FBS and allowed to seed
for 48 h at 37 °C. Media was then replaced for DMEM with 10%
heme-depleted FBS supplemented with 0.5 mMsuccinyl ace-
tone (for SA-treated cells). For untreated cells, media was
changed for DMEM in 10% regular FBS. Both treated and
untreated samples were allowed to incubate at 37 °C for 24 h.
The cells were then collected by scrapping and lysed using 1.5-
mm zirconium beads (Benchmark Scientific). Lysates were
quantified by Bradford assay. In the meantime, hemin-agarose
beads and Sepharose beads were equilibrated three times by
centrifugation with lysis buffer (0.1% Triton X-100, 10 mMso-
dium phosphate, 50 mMKCl, 5 mMEDTA, pH 7.5, 1 3protease
arrest, and RNasin RNase Inhibitor (Promega)). 100 ml of beads
(50-ml bed volume) were used per biological replicate. After
bead equilibration, each lysate was divided into two and 10 mg
were loaded to hemin-agarose and 10 mg to Sepharose beads.
The mixtures were allowed to bind for 60 min, rotating at 20
rpm at room temperature. Then, three washes were performed
using lysis buffer and supernatants were discarded. Each wash
consisted of 10 min of incubation at room temperature with 20
rpm rotation followed by centrifugation at 700 3gfor 5 min.
Bead-bound fractions were eluted by a 15-min incubation at
room temperature with 20-rpm rotation in 50 mlof1Mimida-
zole in lysis buffer, then centrifuged at maximum speed for 2
min, and supernatants were collected. RNA was then extracted
from eluted fractions with TRIzol using the manufacturer’s
protocol. For the PhenDC3 titration in the HEK293 cells
experiment, the same protocol was followed, with the differ-
ence that the cells were allowed to seed for 24 h (20% initial
confluency), and then PhenDC3 was added in increasing con-
centrations (5 mM,10mM,and20mM). DMSO carrier treatment
was performed the same way but with equivalent DMSO vol-
umes. The cells were left at 37 °C for 48 h and collected and
lysed as described above.
RT-qPCR
The sets of primers used can be found in Table S2.LunaUni-
versal One-Step RT-qPCR kit (New England Biolabs) was used
following the manufacturer’s protocol. Fold enrichments were
calculated by comparing the C(t) values obtained from RNAs
extracted from hemin-agarose to RNAs extracted from Sephar-
ose beads. Three biological replicates were performed for all
the RT-qPCR experiments. For BioTASQ experiments, fold
enrichments were calculated by comparing the C(t) values
obtained from the lysates containing BioTASQ 1beads with
those containing beads only.
Heme bioavailability assay using the HS1 sensor
HEK293 cells were plated and transfected in polystyrene-
coated sterile 6-well plates (Greiner) for flow cytometry. The
cells were plated in basal growth medium DMEM containing
10% FBS. At 30% confluency, the cells were transfected with the
heme sensor plasmid pEF52
a
-hHS1 using Lipofectamine LTX
according to the manufacturer’s protocols. After 48 h of treat-
ment with transfection reagents, the cells were treated with
PhenDC3 (1 mMstock) in fresh DMEM 10% FBS for 24 h prior
to harvesting. Heme-depleted cells were treated with 500 mM
SA in DMEM containing 10% heme-depleted FBS for 72 h prior
to harvesting. Heme-sufficient cells were treated with 350 mM
ALA in DMEM 10% FBS for 24 h. The cells were harvested in
13PBS for flow analysis. Flow cytometric measurements were
performed using a BD FACSAria III Cell Sorter equipped with
an argon laser (ex 488 nm) and a yellow-green laser (ex 561
nm). Enhanced GFP (eGFP) was excited using the argon laser
and was measured using a 530/30-nm bandpass filter, and
mKate2 was excited using the yellow-green laser and was meas-
ured using a 610/20-nm bandpass filter. Data evaluation was
conducted using FlowJo v10.4.2 software. Single cells used in
the analysis were selected for by first gating for forward scatter
and side scatter, consistent with intact cells, and then fluores-
cence intensities above background were selected by gating for
cells with mKate2. The fraction of sensor bound to heme may
be quantified according to the following equation (40):
%Bound51003R2Rmin
ðÞ
=Rmax 2Rmin
ðÞ

where Ris the median eGFP/mKate2 fluorescence ratio in reg-
ular media and R
min
and R
max
are the median sensor ratios
when the sensor is depleted of heme or saturated with heme.
R
min
and R
max
values are derived from cells cultured in
HD1SA or in media conditioned with ALA (40). The plot in
Fig. 4Dwas obtained by fitting the median sensor ratios in Fig.
4Cto the following one-site binding model (40,68):
Ratio ¼initial ratio 1Dratio 3ðx=ðKd1xÞÞ
where xis the independent variable, PhenDC3.
Human ribosomes appropriate heme
14862 J. Biol. Chem. (2020) 295(44) 14855–14865
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BG4 purification
pSANG10-3F-BG4 was a gift from Shankar Balasubramanian
(Addgene plasmid 55756; RRID:Addgene_55756). BL21 cells
transformed with this plasmid were grown in room tempera-
ture and induced overnight with 0.1 mMisopropyl 1-thio-
b
-D-
galactopyranoside. The cells were pelleted, then resuspended
in xTractor (Takara Bio) supplemented with Protease Arrest
(G-Biosciences), lysozyme, and DNase I. Sonicated cell lysate was
combined with nickel-nitrilotriacetic acid resin (Invitrogen) and
purified via the His-tag. BG4 was further purified by FPLC using
a Superdex75 size exclusion column (GE Healthcare).
Immunofluorescence
Immunofluorescence was performed by standard protocols.
HEK293 cells were seeded onto poly-L-lysine–coated cover
glass 2 days before the experiment and fixed in 4% formalde-
hyde for 15 min. The cells were permeabilized with 0.1% Triton
X-100 for 3 min and blocked with 5% donkey serum (Jackson
ImmunoResearch Laboratories), followed by incubation with
antibodies for 1 h at room temperature or overnight at 4 °C.
Antibodies used here are BG4, rabbit anti-FLAG (Cell Signaling
Technology, 14793S), mouse anti-L19 (Santa Cruz Biotechnol-
ogy, sc-100830), mouse anti-rRNA (Santa Cruz Biotechnology,
sc-33678), mouse anti-Calnexin (Santa Cruz Biotechnology, sc-
23954), Alexa Fluor 488 conjugated donkey anti-rabbit (Jackson
ImmunoResearch Laboratories, 711-545-152), and Rhodamine
Red-X conjugated donkey anti-mouse (Jackson ImmunoRe-
search Laboratories, 715-295150). After staining, the cells were
carefully washed with Dulbecco’s PBS supplemented with 0.1%
Tween 20. Nuclear DNA was stained with 49,6-diamidino-2-
phenylindole. Images were acquired with a Zeiss 700 Laser
Scanning Confocal Microscope. PhenDC3 treatment consisted
of incubation at 37°C overnight at 10 mM. PhenDC3 treatment
was done prior to cell fixation. Determination of colocalization
ratios was performed as described in Zen software (Zeiss).
No-primary-antibody controls and RNase A–and PhenDC3-
treated images are reported in Fig. S6 and Fig. S7. The colocali-
zation image in Fig. 2, Aand Bshows the G4 signal that colocal-
izes with L19 and with the ER (yellow pixels) and the one that
does not colocalize (green pixels). L19,ER,andBG4 images
only present their respective fluorescence signals.
BioTASQ capture of cellular RNAs
BioTASQ experiments followed published protocols in vitro
(8)andin vivo (22). Briefly, HEK293 cells were seeded onto a 6-
well plate at 20% confluency and allowed to incubate at 37 °C
for 48 h. The cells were then cross-linked with 1% paraformal-
dehyde/PBS for 5 min at room temperature. Cross-linking was
stopped by incubating cells with 0.125 Mglycine for 5 min at
room temperature. The cells were harvested by scrapping and
resuspended in lysis buffer (200 mMKCl, 25 mMTris-HCl, pH
7.5, 5 mMEDTA, 0.5 mMDTT, 1% Triton X-100, RNasin RNase
Inhibitor, and 13protease arrest). The cells were lysed by soni-
cation (30% amplitude, 10 s on and off intervals, and 2 minson-
ication time). The lysate was then split: BioTASQ was added at
a final concentration of 100 mMto one of the samples, and the
other sample was left untreated. The lysates were incubated at
4 °C overnight with gentle rotation. Sera-Mag magnetic
streptavidin-coated beads (GE Healthcare) were washed
three times with wash buffer (5 mMTris-HCl, pH 7.5, 0.5 mM
EDTA, and 1 MKCl). Each wash was followed by centrifuga-
tion at 3,500 rpm for 5 min at 4 °C. The beads were then
treated with buffer 1 (0.1 MNaOH, and 0.05 MKCl in RNase/
DNase-free water) two times at room temperature for 2 min
and then centrifuged at 3,500 rpm at 4 °C for 5 min and
washed with buffer 2 (0.1 MKCl in RNase/DNase-free water).
Lastly, to block, the beads were treated with 1 mg/ml BSA and
1mg/ml yeast tRNA and allowed to incubate at 4 °C overnight
with gentle rotation.
After incubation overnight with BioTASQ, the cell lysates
were treated with 1% BSA for 1 h at 4 °C. Washed magnetic
beads were added to the lysates (20 mg beads/sample) and
allowed to mix with gentle rotation at 4 °C for 1 h. The beads
were then washed three times with lysis buffer for 5 min, and
then cross-linking was reversed by incubating the beads at
70 °C for 1 h. Finally, TRIzol was used to extract RNAs for anal-
ysis by RT-qPCR.
Data availability
All data are contained within this article and in the supporting
information.
Acknowledgments—We thank Drs. Rebecca Donegan, David A.
Hanna, Jonathan B. Chaires, Aaron Engelhart, David Monchaud,
and Judy Wong, and Claudia Montllor-Albalate for helpful discus-
sions. We acknowledge Andrew Shaw and the core facilities at the
Parker H. Petit Institute for Bioengineering and Bioscience at the
Georgia Institute of Technology for expert advice and the use of
equipment. Purified human ribosomes and polysomes were a gift
from Immagina BioTechnology. BioTASQ was a gift from Dr.
David Monchaud.
Author contributions—S. M.-F., A. R. R., and L. D. W. conceptuali-
zation; S. M.-F., C. I., C. M. M., and A. R. R. data curation; S. M.-F.,
and C. I. formal analysis; S. M.-F. and C. I. validation; S. M.-F., C. I.,
and C. M. M. investigation; S. M.-F. and C. I. visualization; S. M.-F.
methodology; S. M.-F., A. R. R., and L. D. W. writing-original draft;
A. R. R. and L. D. W. resources; A. R. R. and L. D. W. supervision;
A. R. R. and L. D. W. funding acquisition; A. R. R. and L. D. W. pro-
ject administration.
Funding and additional information—This work was supported by
NASA Grants 80NSSC17K0295 and 80NSSC18K1139 (Center for
the Origin of Life) (to L. D. W.), the National Institutes of Health
Grant ES025661 (to A. R. R.), and the National Science Foundation
Grant MCB-1552791 (to A. R. R.). The content is solely the respon-
sibility of the authors and does not necessarily represent the official
views of the National Institutes of Health.
Conflict of interest—The authors declare that they have no conflict
of interest with the contents of this article.
Abbreviations—The abbreviations used are: G4, G-quadruplex;
rProtein, ribosomal protein; ER, endoplasmic reticulum; SSU, small
Human ribosomes appropriate heme
J. Biol. Chem. (2020) 295(44) 14855–14865 14863
at University of California, Berkeley on October 30, 2020http://www.jbc.org/Downloaded from

subunit; LSU, large subunit; SA, succinylacetone; HS, heme sensor;
HD1SA, heme-deficient media containing SA; ALA, 5-aminolevu-
linic acid; eGFP, enhanced GFP; ex, excitation; em, emission.
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J. Biol. Chem. (2020) 295(44) 14855–14865 14865
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Williams
Santi Mestre-Fos, Chieri Ito, Courtney M. Moore, Amit R. Reddi and Loren Dean
Human ribosomal G-quadruplexes regulate heme bioavailability
doi: 10.1074/jbc.RA120.014332 originally published online August 13, 2020
2020, 295:14855-14865.J. Biol. Chem.
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