The 30-nm Fiber Redux
Andrew Travers 1, 2
Do alternating stacking modes of nucleosomes
underlie the compaction of chromatin?
The DNA of eukaryotic cells is pack- aged onto nucleosomes—complexes composed of histone proteins—
that together form chromatin, enabling
tight packing of the genome within the cell
nucleus. In the first folded structure of chro-
matin to be characterized, nucleosomes were
coiled into a ~30-nm-diameter helix, with
the “linker” histone located in the interior of
the fiber (1). The precise molecular organiza-
tion of this 30-nm fiber has long been exten-
sively debated. Initially, structural studies on
fibers assembled on natural DNA sequences
were hampered by variation in the length of
the linker DNA between nucleosomes. How-
ever, more recently, the construction of reg-
ularly spaced tandem DNA repeats for pre-
cise nucleosome positioning (2) has revolu-
tionized analysis. On page 376 of this issue,
Song et al. (3) determine by cryo–electron
microscopy the 11 Å–resolution structure
of 30-nm fibers assembled from arrays of
Song et al. unequivocally identify the path
of linker DNA. They show that in the fiber, a
linear array of nucleosomes is packed in two
interwound left-handed helical stacks with
a straight linker DNA between successive
nucleosomes in the array crossing the interior
of the fiber. In agreement with an emerging
consensus, this finding resolves the funda-
mental issue as to whether the fiber is built
from one nucleosome stack—a solenoid—
or from two. However, importantly, the new
structure differs in one largely unanticipated
aspect from most previous models. Instead of
the monotonous helix previously imposed by
the limitations of the available information,
the 30-nm fiber is formed by the tight heli-
cal packing of a tetranucleosome unit. Within
this unit, first observed by the crystallization
of a tetranucleosome lacking linker histone
(4), the two opposing nucleosome dimers are
fully stacked on each other with only a small
angular separation. Between the units, the
angular separation is larger and each unit is
staggered relative to its neighbor. The struc-
ture agrees well with all other direct mea-
surements but does not exclude the possibil-
ity that a nucleosome array may still have the
potential to fold into other helical forms.
The first hint of a lack of structural uniformity of nucleosomes in the folded fiber came
from the observation of alternating pattern
of deoxyribonuclease I (DNase I) digestion
in successive nucleosomes (5). Subsequent
modeling of the fiber showed that retention
of the tetranucleosome unit necessitated the
1MRC Laboratory of Molecular Biology, Francis Crick Avenue,
Cambridge CB2 0QH, UK. 2Department of Biochemistry,
University of Cambridge, 80 Tennis Court Road, Cambridge
CB2 1GA, UK. E-mail: email@example.com
an action potential. However, light-sensitive
ion channels for silencing neurons must be
selective for chloride (Cl–) or potassium (K+)
to further decrease the membrane potential
(hyperpolarization) and thereby turn off the
action potential. Such ion channels have not
yet been found in natural species. In optogenetics, light-driven chloride and proton
pumps from archaeal halobacteria have been
used instead; however, pumps conduct only
a single ion per photon and are therefore less
efficient than channels.
Wietek et al. and Berndt et al. now provide drastic solutions to this problem. They
use molecular engineering to convert cation-conducting channelrhodopsins into light-gated ion channels that selectively conduct
Cl–. Further, they show that these engineered
opsins can be used to efficiently hyperpolarize the membrane.
The conversions of ion selectivity
obtained by the two groups occur in com-
pletely different ways (see the figure).
Wietek et al. convert the ion selectivity of
their channel through a single mutation,
replacing an acidic residue with a positively
charged arginine at a gating site located in
the middle of the putative pore (see the fig-
ure, panel B). This observation suggests
that the gating site may act as a selectivity
filter. In contrast, Berndt et al. achieve Cl−
selectivity through multiple mutations that
alter the electrostatic environment along an
extended region of the pore, while replacing
the acidic residue at the gating site by a neu-
tral one (see the figure, panel C). The mech-
anisms underlying the Cl− selectivity of the
two engineered channels are thus remark-
Ion channels are thought to achieve high
ion selectivity through precisely defined
pore architectures that efficiently conduct
particular ions, as seen in tetrameric K+
selective channels (8). However, the different ion selectivity mechanisms of the two
engineered channelrhodopsins show that
charge selectivity of ion channels can be
controlled in more variable ways. A clue to
the permissive feature of channelrhodopsins
may come from experimental evidence that
channelrhodopsin is a light-driven proton
pump as well as an ion channel (9) and that
a single mutation in the middle of the proton channel can convert another light-driven
opsin proton pump into a Cl– pump (10).
In both systems, the proton pump function
is fulfilled by concerted motion of protons
between acidic residues, positively charged
basic side chains, and water molecules in
the pore (11); the flexibility of these residues may contribute to making channelrhodopsins amenable to molecular engineering.
The highly controllable charge selectivity of engineered channelrhodopsins demonstrated by Wietek et al. and Berndt et al.
should encourage researchers to further augment and improve functionalities of channelrhodopsins for optogenetics applications. Understanding of the physicochemical mechanism by which channelrhodopsins
operate will also help to guide the design of
novel ion channels with controllable selectivity through de novo protein design (12),
synthetic engineering (13), and biomimetic
synthesis of artificial macromolecules.
1. K. Deisseroth, Sci. Am. 303, 48 (2010).
2. F. Zhang et al., Cell 147, 1446 (2011).
3. J. Wietek et al., Science 344, 409 (2014);
4. A. Berndt, S. Y. Lee, C. Ramakrishnan, K. Deisseroth,
Science 344, 420 (2014).
5. H. E. Kato et al., Nature 482, 369 (2012).
6. G. Nagel et al., Science 296, 2395 (2002).
7. G. Nagel et al., Proc. Natl. Acad. Sci. U.S.A. 100, 13940
8. D. A. Doyle et al., Science 280, 69 (1998).
9. K. Feldbauer et al., Proc. Natl. Acad. Sci. U. S.A. 106,
10. J. Sasaki et al., Science 269, 73 (1995).
11. O. P. Ernst et al., Chem. Rev. 114, 126 (2014).
12. N. R. Zaccai et al., Nat. Chem. Biol. 7, 935 (2011).
13. W. Grosse, L.-O. Essen, U. Koert, ChemBioChem 12, 830