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The Journal of Neuroscience, May 15, 1998, 18(10):3738-3748
Two Phases of Rod Photoreceptor Differentiation during Rat
Retinal Development
Eric M.
Morrow,
Michael J.
Belliveau, and
Constance L.
Cepko
Department of Genetics and Howard Hughes Medical Institute, Harvard
Medical School, Boston, Massachusetts 02115
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ABSTRACT |
We have conducted a comprehensive analysis of the relative timing
of the terminal mitosis and the onset of rhodopsin expression in rod
precursors in the rat retina in vivo. This analysis
demonstrated that there are two distinct phases of rod development
during retinal histogenesis. For the majority of rod precursors, those
born on or after embryonic day 19 (E19), the onset of rhodopsin
expression was strongly correlated temporally with cell cycle
withdrawal. For these precursors, the lag between the terminal mitosis
and rhodopsin expression was measured to be 5.5-6.5 d on average. By
contrast, for rod precursors born before E19, the lag was measured to
be significantly longer, averaging from 8.5 to 12.5 d. In
addition, these early-born rod precursors seemed to initiate rhodopsin
expression in a manner that was not correlated temporally with the
terminal mitosis. In these cells, onset of rhodopsin expression
appeared approximately synchronous with later-born cells, suggesting a synchronous recruitment to the rod cell fate induced by environmental signals. To examine this possibility, experiments in which the early-born precursors were exposed to a late environment were conducted, using a reaggregate culture system. In these experiments, the early-born precursors appeared remarkably uninfluenced by the late
environment with respect to both rod determination and the kinetics of
rhodopsin expression. These results support the idea that intrinsically
distinct populations of rod precursors constitute the two phases of rod
development and that the behavior exhibited by the early-born
precursors is intrinsically programmed.
Key words:
rodent retina; rod photoreceptors; rhodopsin; cell fate
determination; differentiation; neurodevelopment
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INTRODUCTION |
Rods are highly specialized,
light-sensing neurons with elaborate outer segments containing the
photopigment rhodopsin. Although the morphology and molecular
physiology of rod photoreceptors have been well studied, the steps that
lead from a multipotent retinal progenitor to a committed rod precursor
and finally to a terminally differentiated rod remain poorly
understood.
Retinal cell-type specification is likely to involve an interaction
between extrinsic and intrinsic regulators (for review, see Reh, 1991 ;
Cepko et al., 1996). Extrinsic cues implicated in rod development
in vitro include taurine (Altshuler et al., 1993), S-laminin
(Hunter et al., 1992), retinoic acid (Kelley et al., 1994), and the
CNTF family of cytokines (Fuhrmann et al., 1995; Kirsch et al., 1996;
Ezzeddine et al., 1997; Neophytou et al., 1997). These studies have all
used rhodopsin expression as an assay for rod differentiation. To date,
two transcription factors have been molecularly characterized that bind
to and transactivate from promoters of photoreceptor-specific genes
in vitro, namely NRL (Kumar et al., 1996; Rehemtulla et al.,
1996) and Crx (Chen et al., 1997; Furukawa et al., 1997). The roles
that the above-mentioned cell autonomous and cell nonautonomous
activities play in rod commitment, and the potential timing of action
of these factors in the life history of a developing rod
in vivo, are areas of active study.
Lineage analysis supports the idea that commitment to the rod cell fate
may occur during or after the terminal mitosis because two cell clones
with one rod and a second distinct cell type were found (Turner and
Cepko, 1987). As well, other studies have directly demonstrated
persistent developmental plasticity in some postmitotic retinal cells
in vitro (Belecky-Adams et al., 1996; Ezzeddine et al.,
1997). In one of these studies, in which developing retinal explants
were treated with ciliary neurotrophic factor, postmitotic rod
precursors were respecified to express at least three markers of
bipolar interneurons, and rhodopsin expression was blocked, suggesting
a switch in cell fate. Furthermore, the window of sensitivity to the
effects of factor treatment in rod precursors was found to extend to
shortly before the expression of rhodopsin (Ezzeddine et al.,
1997).
Several studies across different species have reported a long delay
between the terminal mitosis of a rod precursor and the onset of
rhodopsin expression (for review, see Cepko, 1996). In the present
study, we set out to answer the following question in the developing
rat retina: is there a correlation between the day a rod precursor is
born and the day it expresses rhodopsin? Our results demonstrate that
there is a strong temporal correlation between the day of birth of a
rod precursor and the onset of rhodopsin expression among cells born on
or after embryonic day 19 (E19). Interestingly, a different behavior is
exhibited by rod precursors born before E19, indicating that there are
two phases of rod development: an early phase and a late phase. We
tested whether extrinsic cues from the late phase could alter the
differentiation of rods born during the early phase. The results are
consistent with the notion that intrinsically distinct populations of
rod precursors participate in the two different phases of rod
development.
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MATERIALS AND METHODS |
Animals. Timed-pregnant Sprague Dawley rats were
purchased from Taconic (Germantown, NY). Most of the litters were born
on E22, which was considered equivalent to postnatal day 0 (P0).
In vivo [3H]thymidine
pulse-labeling. [3H]thymidine (Amersham,
Arlington Heights, IL) (5 µCi/gm of animal) was administered by a
single intraperitoneal injection to pregnant rats (embryonic time
points) or separately to neonatal littermates (postnatal time
points).
In vitro [3H]thymidine pulse-labeling
and BrdU labeling of retinal explants. Pulse-labeling in
vitro was performed by placing P0 explanted retinae in DMEM (Life
Technologies, Gaithersburg, MD) containing 10% fetal calf serum (FCS)
(Life Technologies) to which a final concentration of 5 µCi/ml
[3H]thymidine was added. After a 1 hr incubation
at 37°C, retinae were washed five times and cultured as explants as
described previously (Lillien and Cepko, 1992) in DMEM containing 10%
FCS and penicillin-streptomycin (100 U/ml) (Life Technologies). Cells
undergoing S-phase in culture were labeled by BrdU addition after
different time intervals to a final concentration of 10 mM.
Retinal dissociations. A developmental series (P0-P20) of
retinae from [3H]thymidine-labeled littermates or
tissue from cultured explants was analyzed by gentle dissociation.
Dissociations were conducted as described by Altshuler and Cepko (1992)
with slight modification. Briefly, neural retinae were dissected free
of other ocular tissues and incubated for 10 min at room temperature in
HBSS lacking Ca2+/Mg2+ (Life
Technologies) to which trypsin (Worthington, Freehold, NJ) was added to
a final concentration of 1 mg/mL. After trypsinization, soybean trypsin
inhibitor (Sigma, St. Louis, MO) was added to a final concentration of
2 mg/ml. The cells were then pelleted by centrifugation (1200 rpm, 5 min), resuspended, and gently triturated to a single cell suspension in
HBSS containing 100 µg/ml DNase I (Sigma). Cells were then plated on
poly-D-lysine (Sigma)-coated, eight-well glass slides
(Cel-Line Associates, Newfield, NJ) before fixation.
Immunocytochemistry and autoradiography. Slides with
[3H]thymidine-labeled cells were first
immunostained and then processed for autoradiography. After they were
plated, retinal cells were fixed with 4% paraformaldehyde for 5 min
before they were blocked for 30 min in PBS, 2% donkey serum (Jackson
ImmunoResearch, West Grove, PA), and 0.1% Triton X-100 (Sigma). The
Rho4D2 monoclonal antibody raised against bovine rhodopsin was a
generous gift of Dr. R. S. Molday (University of British Columbia)
and was used at a 1:250 dilution (Molday, 1989). BrdU detection was
performed using the anti-BrdU monoclonal antibody containing nuclease
(Amersham) as instructed by the supplier, with the following
modification. Cells were treated with 2N HCl for 20 min after fixation,
followed by eight washes with HBSS, pH 7.4, before they were blocked. A Texas Red-conjugated, donkey anti-mouse IgG secondary antibody (Jackson
ImmunoResearch) was used according to the supplier's instructions for
indirect immunodetection. Nuclear staining was performed by adding
4',6-diamidine-2-phenylindole-dihydrochloride (DAPI) to the secondary
antibody solution to a final concentration of 0.0005%. After they were
immunostained, the slides were dehydrated and stored at 4°C before
autoradiography.
Dehydrated slides with [3H]thymidine-labeled cells
were dipped in NTB2 autoradiography emulsion (Kodak, Rochester, NY).
Slides were stored at 4°C in the dark while they were exposed for
4-10 weeks for in vivo labeling experiments or 1-2 weeks
for in vitro labeling experiments. Slides were then
developed for 5 min in D19 developer (Kodak) and rinsed in distilled
water followed by 20 min in fixer (Kodak). Slides were washed with
distilled water for 10 min and then mounted in gelvatol (Rodriquez and
Dunhardt, 1960).
Fluorescent microscopy and scoring cells undergoing terminal
mitosis. Doubly processed slides were viewed using a Zeiss
Axiophot fluorescent microscope. Silver grains were counted under
transmission light microscopy using a 63× Plan NEOFLUAR objective
(Zeiss). Cells were determined to have undergone their terminal mitosis on the day of the [3H]thymidine pulse if they
contained more than half the number of silver grains of the most
heavily labeled cell in a given sample.
Reaggregate pellet cultures. The reaggregate pellet culture
protocol was modified from Watanabe and Raff (1990) (see schematization in Fig. 6A). Retinae from E16 rats were dissected and
labeled with 5 µCi/ml [3H]thymidine for 1 hr
before dissociation (as described above). [3H]thymidine-labeled E16 retinae were
dissociated, and cells were counted and pelleted in a microcentrifuge
tube containing 20- to 50-fold excess unlabeled P0 retinal cells by
centrifugation for 7 min at 1150 × g. The total number
of cells per pellet was 5 × 105 cells. Pellets
were transferred to nucleopore polycarbonate membranes, 0.2 µm pore
size (Costar Nucleopore, Charlotte, NC), and cultured for 3-17 d as
described for explants (Lillien and Cepko, 1992) in 45% DMEM, 45%
Ham's F12 Nutrient Mixture (Life Technologies), 10% FCS, and
penicillin-streptomycin (100 U/ml). At the end of the culture period
pellets were dislodged from the membranes, dissociated, and processed
autoradiographically and immunocytochemically as described above. PKH-2
and PHK-26 dye-labeled cells were produced according to the directions
of the supplier (Sigma). Cells were labeled in the indicated dye
diluted to 4 µM in diluent for 5 min. The labeling
reaction was halted by adding an equal volume of FCS and then washed
several times in culture medium. Reaggregate cultures were produced as
described above. Imaging of dye-labeled cells was performed on a Leica
TCS-NT confocal microscope.
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RESULTS |
Measuring the timing of the terminal S-phase in retinal cells
A goal of this study was to conduct a comprehensive analysis of
the relative timing of cell cycle withdrawal and the onset of rhodopsin
expression in developing rods in vivo. To measure the timing
of cell cycle withdrawal of retinal cells in vivo, the
classic [3H]thymidine "birthdating" method was
used (Sidman, 1970). This method involves administering a pulse of
[3H]thymidine in vivo during
development. Cells in S-phase at the time of the pulse incorporate
[3H]thymidine into the DNA of their daughter
cells. Daughter cells that continue to proliferate subsequent to the
pulse dilute the [3H]thymidine in an approximately
binary manner with each cell division. Daughters of cells undergoing
their terminal S-phase at the time of the pulse remain "heavily
labeled" and are termed "born" at the time of or shortly after
the pulse. The degree of [3H]thymidine labeling
can be measured by scoring silver grain numbers after autoradiographic
processing of the tissue.
For the present study, two modifications to the classic
[3H]thymidine birthdating procedure were made.
First, to facilitate cell and silver grain counting, heavily labeled
cells were scored after dissociation of the retina. This was found to
be necessary for accurate quantification because of high cell density
in the outer nuclear layer and the small size of rod nuclei. The second modification to the classic birthdating method was that instead of
relying on morphology to identify cell type, we used indirect immunocytochemistry against rhodopsin protein using the anti-rhodopsin antibody Rho4D2 (Molday, 1989). Figure
1A,B shows an example
of a heavily labeled cell that is immunoreactive for rhodopsin.
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Figure 1.
Immunofluorescent and autoradiographic analysis of
dissociated retinal cells. A, Cell heavily labeled for
[3H]thymidine that was also immunoreactive with
Rho4D2, shown in B (large arrow).
C, Cell heavily labeled for
[3H]thymidine that was also immunoreactive for
BrdU incorporation, shown in D (large
arrow). Arrowheads mark examples of
immunoreactive cells that were not heavily labeled by
[3H]thymidine. Small arrows mark
examples of cells that were negative for staining with the
corresponding primary antibody. Cells in A and
C were viewed under bright-field and UV optics. The
nuclei were stained by the nuclear stain DAPI. Cells in
B and D were viewed under fluorescent
light conditions suitable for detecting the Texas Red-conjugated
secondary. Note that Rho4D2 staining was membrane-associated, whereas
staining for BrdU incorporation was nuclear.
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In addition to these modifications, the following control studies were
performed to evaluate the criterion for heavily labeled cells. The
classic studies (Carter-Dawson and LaVail, 1979) and other more recent
studies (Alexiades and Cepko, 1997; Ezzeddine et al., 1997) have used
more than half the grain number of the most heavily labeled cell in an
experiment as the criterion for heavily labeled. Another recent study
has used the criterion of one-fourth or greater (Arimatsu et al.,
1994). Figure 2A
presents examples of distributions of grain number for retinae injected on E17 and harvested on P1 or P15. To test the cutoff of more than half
the most heavily labeled cell, cells were pulse-labeled in
vivo at P0 and in vitro in P0 explant culture.
Subsequent to [3H]thymidine pulse-labeling,
littermates were pulsed with BrdU, or BrdU was added to a retinal
explant culture for cumulative labeling, at 5, 12, 24, or 48 hr after
[3H]thymidine pulse. At P4 in vivo or
after 4 d of culture, the percentage of
[3H]thymidine heavily labeled cells that
incorporated BrdU at the different times of BrdU addition was
evaluated. In this way, the length of time after the
[3H]thymidine pulse when heavily labeled cells
(using the criterion of more than half the most heavily labeled cell)
stopped incorporating BrdU, and therefore left the cell cycle, could be
evaluated. Figure 1C,D shows an example of a
[3H]thymidine heavily labeled cell that was also
positive for BrdU incorporation.
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Figure 2.
[3H]thymidine birthdating as
a method for labeling retinal cells undergoing their terminal S-phase.
A, Representative histograms displaying grain
distributions in cells of retinae that were administered a pulse of
[3H]thymidine at E17 in vivo and
harvested at P1 or P15. The criterion for "heavily labeled" (as
indicated) is defined as cells containing more than half the grain
number of the most heavily labeled cell in an experiment. In the
supporting table, the number of heavily labeled cells
that survived to P1 and P15 was approximated. B,
Cumulative BrdU labeling after [3H]thymidine pulse
in vitro. P0 retinal explant cultures were administered
a 1 hr pulse of [3H]thymidine in
vitro and then cumulatively labeled by addition of BrdU to
cultures at 5, 12, 24, or 48 hr after the
[3H]thymidine pulse. Retinae were then examined
after 4 d of culture for doubly labeled cells. Plotted are the
percentages of [3H]thymidine heavily labeled cells
that were positive for BrdU incorporation at the different times of
addition. Each data point represents the average ± SEM
(n = 4). More than 50 heavily labeled cells were
scored per trial for each time point.
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Figure 2B shows the results from the in
vitro explant control experiment. These data show that 91.5 ± 1.0% of cells were heavily labeled with
[3H]thymidine and labeled for BrdU incorporation
when BrdU was added to the culture 1 hr after the
[3H]thymidine pulse, or 49.4 ± 2.2% when
the BrdU was added 5 hr after the [3H]thymidine
pulse. However, when BrdU was added 12, 24, or 48 hr after the
[3H]thymidine pulse, only 11.3 ± 3.7, 8.0 ± 1.8, or 3.0 ± 1.9%, respectively, of cells were
doubly positive. These data are consistent with the S-phase length of
18 hr at P0, as measured previously (Alexiades and Cepko, 1996).
In vivo, 0% (0/155 and 0/101; n = 2) of
[3H]thymidine heavily labeled cells were labeled
by BrdU administration 48 hr after the
[3H]thymidine pulse. From these data, we conclude
that, by the described criterion for heavily labeled, >92% of heavily
labeled cells have undergone their terminal S-phase by 24 hr after the
[3H]thymidine pulse, and >97% by 48 hr.
Finally, the number of cells born on E17 in vivo that
survived to P1 and P15 was approximated to be 2.69 × 105 and 2.11 × 105,
respectively (see table in Fig. 2A). On the basis of
these numbers and the number of postmitotic cells expected to be
generated on E17, 3.21 × 105 (Alexiades and
Cepko, 1996), the percentage of cells surviving from E17 to P1 and from
P1 to P15 was estimated to be 83% (2.69 × 105
divided by 3.21 × 105) and 78% (2.11 × 105 divided by 2.69 × 105),
respectively. The cell loss seen was therefore consistent with two
reports on normal cell death (Young, 1984; Alexiades and Cepko, 1997).
A third study on cell death in this period may have predicted a greater
cell loss between P1 and P15 (Voyvodic et al., 1995). Regardless of
this discrepancy, these data support the interpretation that the
[3H]thymidine administration was nontoxic to the
developing retina. Similar analyses of the results from other injection
time points were also consistent with this interpretation (data not
shown).
Long average latency (6.5-7.0 d) between terminal S-phase and
onset of rhodopsin expression in vivo
The [3H]thymidine birthdating method was
first used to measure the average lag between cell cycle withdrawal and
the onset of rhodopsin expression for the total population of
developing rods. To this end, the kinetics of cell cycle withdrawal and
the onset of rhodopsin expression were initially analyzed independently for the population of developing rods. These measurements were then
correlated to derive an average lag.
First, the [3H]thymidine birthdating method was
applied to evaluate the overall kinetics of rod precursor genesis.
Pregnant rats and neonatal littermates were injected with
[3H]thymidine, and the percentage of surviving
cells born on the different injection days that ultimately
differentiated as rods was determined using combined
autoradiographic-immunocytochemical analysis on mature retinae. As
shown in Figure 3A, rod
precursors were generated as early as E14, when 5.0 ± 0.4% of
surviving cells differentiated as rods. The peak generation of rod
precursors occurred between E21 and P2, when 78.3 ± 8.0 and
79.0 ± 4.5%, respectively, of surviving cells born on these days
differentiated as rods. Rod precursor genesis continued into the late
stages of the first postnatal week, when cell proliferation ceased
entirely (Alexiades and Cepko, 1996).
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Figure 3.
Overall kinetics of rod precursor genesis
correlated with kinetics of rhodopsin onset. A, Rod
precursor genesis in developing rat retina. Developing retinae were
administered a [3H]thymidine pulse in
vivo on E14, E15, E17, E19, E21, P0, P2, or P5. The percentage
of surviving cells born on each day of injection that differentiated as
rod photoreceptors was assessed using combined
immunocytochemical-autoradiographic analysis on mature retinae.
B, Kinetics of rhodopsin expression in neonatal retina
and kinetics of onset of immunocytochemically detectable rhodopsin
expression in the postnatal rat retina. Plotted are the percentages of
cells per retina positive for Rho4D2 staining during postnatal
development. C, The kinetics of rhodopsin onset closely
mimic the kinetics of rod precursor genesis separated by a 6.5-7.0 d
lag. We used the data in A and B and the
number of total retinal cells and postmitotic cells generated on each
day of retinal development (Alexiades and Cepko, 1996) to approximate
the cumulative numbers of rod precursors born and rhodopsin-positive
cells for each day of retinal development.
(The estimation of the cumulative number of
rod precursors born neglects cell death and therefore may be a slight
overestimate for the earlier time points in particular.) The
plot lines for A and for rod precursor
birth in C represent the average of two trials, and the
error range extends to the values of each individual trial, with the
exception of P5 where a single trial was performed. One hundred or more
heavily labeled cells were scored for each data point for these curves.
In B and for the number of rhodopsin-positive cells in
C, each plotted value represents the average ± SEM.
More than 250 cells per trial were scored, with between two and six
trials per time point.
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The kinetics of onset of rhodopsin expression was analyzed over the
full course of postnatal retinal development in vivo. To
this end, a developmental series of freshly dissociated retinae was
examined using the anti-rhodopsin antibody Rho4D2. Because rhodopsin
protein expression closely mimics rhodopsin gene transcription (Treisman et al., 1988), measurements of the onset of rhodopsin immunoreactivity are likely to reflect upregulation of gene
transcription. The kinetics of onset of immunocytochemically detectable
rhodopsin in the postnatal retina are shown in Figure 3B.
Interestingly, the onset of rhodopsin expression was found to be
roughly biphasic. The percentage of rhodopsin-positive cells increased
slowly from <1% at P1 to 10.5 ± 0.8% at P6 and then increased
rapidly to 68.8 ± 2.9% by P10. Ultimately, 71.6 ± 2.5% of
retinal cells were rod photoreceptors in the mature retina.
We used the percentage of rods born (Fig. 3A) and the number of cells
born on each day of development (Alexiades and Cepko, 1996) to
approximate the cumulative number of rod precursors born (Fig.
3C). Also, by multiplying the percentage of
rhodopsin-positive cells (Fig. 3B) and the total number of
retinal cells on each day of development (Alexiades and Cepko, 1996),
we calculated the number of rhodopsin-positive cells per retina for
each day of development (Fig. 3C). As shown in Figure
3C, the kinetics of rhodopsin onset closely mimics the
kinetics of rod precursor genesis, separated by a 6-7 d lag. These
data, summarized in Table 1, indicate
that on the basis of an average for the total population of developing
rods, rod precursors display a lag of ~6.5-7.0 d between their
terminal S-phase and the onset of rhodopsin expression.
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Table 1.
The average lag between terminal S-phase and onset of
rhodopsin is 6.5-7.0 d for the total population of developing rods
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Rhodopsin onset is correlated temporally with terminal mitosis in
postnatally born rod precursors
We next set out to determine whether the duration of the latency
between cell cycle withdrawal and rhodopsin onset was fixed or varied
depending on the birthdate of the rod precursor. To investigate this
question for postnatally born rods, the following experiment was
performed. Neonatal littermates were administered single injections of
[3H]thymidine on P0, P2, or P5 to label retinal
precursor cells undergoing terminal S-phases on these days. A
developmental series of freshly dissociated retinae from these litters
was then analyzed autoradiographically and for anti-rhodopsin
immunoreactivity. The kinetics of onset of rhodopsin expression for
cohorts of retinal cells born on each given day was thereby determined
(Fig. 4A-C). Figure
4D illustrates the kinetics of rhodopsin expression
normalized for the percentages of rods ultimately found in each mature
cohort.
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Figure 4.
Kinetics of onset of rhodopsin expression in
retinal cohorts born postnatally. Neonatal litters were administered
[3H]thymidine by intraperitoneal injection on P0,
P2, or P5 to label retinal precursor cells undergoing terminal S-phases
on these days. A developmental series of freshly dissociated retinae
from these litters was then analyzed auto-radiographically and for anti-rhodopsin
immunoreactivity. A, P0.75; B, P2.5;
C, P5.75. D, Co-plot of kinetics for
P0.75, P2.5, and P5.75, normalized for the percentages of rods
ultimately found in each mature cohort. The plot line
represents the average of two trials for each time point. For
A and B, the error range extends to the
values of each individual trial.
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Despite limited asynchrony within cohorts, rhodopsin
onset in rod precursors born postnatally appeared to be strongly
correlated temporally with the terminal mitosis. A fixed lag time for
these cohorts was clearly demonstrated by comparing the lag to 50%
final rhodopsin levels across postnatal cohorts: 5.65 ± 0.27, 5.97 ± 0.01, and 5.72 d for P0-, P2-, and P5-born cohorts,
respectively. Table 2 summarizes the lag
times to 10, 50, and 90% final rhodopsin levels for the postnatal
cohorts.
Early embryonically born rods show a variable lag and express
rhodopsin synchronously with later-born cohorts
Although rod precursors are born as early as E14, the number of
immunocytochemically detectable rhodopsin-positive cells is very low in
the early neonatal rat retina (Fig. 3). We therefore sought to
characterize the relative timing of the terminal mitosis and the onset
of rhodopsin expression in embryonically born rod precursors in
vivo. To investigate this question, pregnant rats were
administered single injections of [3H]thymidine on
E15, E17, E19, or E21. A developmental series of freshly dissociated
retinae from these litters was then analyzed as done previously for the
postnatally born cohorts. Figure 5 illustrates the results of this experiment for the E17, E19, and E21
cohorts. Data for the E15 cohort is presented in Table
3. As shown in Figure 3A,
10.5 ± 0.5, 16.9 ± 0.1, 65.0 ± 8.0, and 78.3 ± 2.2% of surviving cells born on E15, E17, E19, and E21, respectively,
were fated to differentiate as rods.
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Figure 5.
Kinetics of onset of rhodopsin expression in
retinal cohorts born embryonically. Pregnant rats were administered
single injections of [3H]thymidine on E17, E19,
and E21 to label retinal precursor cells undergoing terminal S-phases
on these days. A developmental series of freshly dissociated retinae
from these litters was then analyzed auto-radiographically and for anti-rhodopsin
immunoreactivity. A, E17.5; B, E19.5;
C, E21.75. D, Co-plot of kinetics for
E17.5, E19.5, and E21.75, normalized for the percentages of rods
ultimately found in each mature cohort. The plot line
represents the average of two trials for each time point. For
A-C, the error range extends to the
values of each individual trial.
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As shown in Figure 5D and summarized in Table
4, the E19 and E21 cohorts displayed
kinetics of rhodopsin onset similar to that of the postnatal cohorts.
The E19 and E21 cohorts displayed a lag of ~6.10 ± 0.07 and
6.54 ± 0.00 d, respectively, to 50% of final levels of
rhodopsin. The E15 and E17 cohorts, however, appeared to display
different kinetics. For the E17 cohort, the lag to 50% of the final
level of rhodopsin-positive cells was 8.24 ± 0.30 d, longer
than the lag of 5.5 to 6.5 d for the later-born rod precursors.
For the E17-born rods, 50% of the final level of rhodopsin-positive
cells were detected on P3.74 ± 0.30, which was synchronous with
the E19 cohort that was born 2 d later and reached 50% final
levels at P3.60 ± 0.07 (Fig. 5D). The E15 cohort had
an even longer lag than the E17 cohort, averaging ~12.5 d (Tables 3,
4). Similarly, for the E15-born cohort, rhodopsin onset also appeared
approximately synchronous with later-born rod precursors. The lag times
to 10, 50, and 90% final rhodopsin levels are summarized for the
embryonically born cohorts in Table 4.
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Table 4.
Timing of rhodopsin onset in early embryonic-born precursor
appears approximately synchronous with later-born cohorts
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Culturing E16-born cells with excess P0 cells does not alter lag to
rhodopsin or change percentage of E16 cells differentiating as rods
As described above, early embryonically born precursors displayed
a long lag to rhodopsin onset and a low frequency of rod differentiation relative to postnatally born precursors. To examine whether this is intrinsically programmed or whether it can be influenced by environmental factors, we performed experiments similar
to those conducted by Watanabe and Raff (1990). As outlined in Figure
6A, intact E16 retinal
explants were pulse-labeled with [3H]thymidine
in vitro and then dissociated and reaggregated with a
20-fold excess of unlabeled E16 retinal cells or P0 retinal cells. The
reaggregate pellets were cultured for 3, 5, 7, 10, 12, 15, or 17 d, at which point rod differentiation was completed. At each of these
time points, the heavily labeled cells, or cells in their terminal
S-phase in the explant, were scored for rhodopsin expression.
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Figure 6.
Culturing embryonically born cells with excess
postnatal cells does not alter lag to rhodopsin or change percentage of
embryonic cells differentiating as rods. A, Outline of
protocol for mixed reaggregate experiments (see Materials and Methods).
B, Kinetics of rhodopsin expression in reaggregates of
E16-born cells cultured alone ( ), P0-born cells cultured alone
( ), or E16-born cells cultured with a 20-fold excess of P0 retinal
cells ( ). C, D, Confocal images of reaggregates
cultured for 1 hr (C) or 5 d
(D). E16 cells are labeled with PKH-26 dye and
are depicted in red; P0 cells are labeled with PKH-2 dye
and are depicted in green. C is one
optical slice, and D is a composite of 16 optical slices
covering 10 µM in the z-axis.
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When [3H]thymidine-labeled E16 cells were
reaggregated with other E16 cells, 0.00 ± 0.00% (0/500) of cells
born in the E16 explant expressed rhodopsin after 5 d (Fig.
6B). The first cells born on E16 that expressed
detectable rhodopsin were observed after 10 d in vitro.
After rhodopsin expression had reached a plateau at 17 d in
vitro, 21.67 ± 4.08% of the cells born on E16 expressed
rhodopsin. For cells born on P0 and reaggregated with other P0 cells,
the first rhodopsin-positive cells were present after 3 d in
vitro. Of such cells, 25.13 ± 5.92% expressed rhodopsin by
5 d in culture, and 67.00 ± 3.65% of such cells expressed
rhodopsin by 17 d in culture (Fig. 6B). The
kinetics and plateau levels of rhodopsin expression roughly mimicked
those observed for cells in vivo as presented above.
When cells born on E16 cells were reaggregated with an excess of
unlabeled P0 cells, the percentage of birthdated E16 cells expressing
rhodopsin was largely unaltered relative to controls at all time points
examined. As shown, 0.00 ± 0.00% (0/500) of cells initiating
their terminal S-phase in E16 explants expressed rhodopsin after 5 d in vitro with excess P0 cells, compared with 0.00 ± 0.00% for such cells cultured with an excess of other E16 cells (Fig.
6B). Again, the first cells born on E16 to become rhodopsin-positive were not detected until 10 d in
vitro, despite the fact that cells from the P0 retina were
beginning to express rhodopsin before this point (data not shown).
After rod differentiation was completed in culture at 17 d
in vitro, 23.33 ± 3.52% of cells born on E16
expressed rhodopsin when cultured with P0 cells versus 21.67 ± 4.08% when cultured with other E16 cells. To exclude the possibility
that a 20-fold excess of P0 cells was not sufficient to influence the
E16 cells, E16 retinal explants were pulse-labeled with
[3H]thymidine in vitro and then
dissociated and reaggregated with a 50-fold excess of unlabeled P0
cells. In such reaggregates, 17.20% (16/93) of the E16 birthdated
cells expressed rhodopsin after 17 d in vitro. The
different behavior exhibited by the E16 and P0 cells in
vitro was not caused by the separation of the two populations in
the pellets. The E16 cells remained well distributed during the
formation of the reaggregate (Fig. 6C) and remained distributed during the culture period (Fig. 6D).
Moreover, within the mixed pellets, birthdated P0 cells expressed
rhodopsin with kinetics similar to that seen for P0 cells in P0
reaggregates (data not shown). We conclude, therefore, that cells born
on E16 were not influenced by the environmental signals created by an excess of P0 cells with respect to both the onset of rhodopsin expression and the percentage of such cells differentiating as rods.
| |
DISCUSSION |
We have conducted a comprehensive analysis of the relative timing
of the terminal mitosis and the onset of rhodopsin expression in the
rat retina in vivo. Our results demonstrate that there are
two distinct phases of rod development during the histogenesis of the
rat retina in vivo: an early phase and a late phase. Figure 7 summarizes our interpretation of the
results from the in vivo analysis. Most rod precursors
participate in the late phase of rod genesis, are born after E19, and
display remarkable regularity in the onset of rhodopsin expression.
These rod precursors initiate rhodopsin expression 5.5-6.5 d on
average after their terminal mitosis. By contrast, for early-phase rod
precursors, i.e., those born before E19, the lag between the terminal
mitosis and the onset of rhodopsin expression was measured to be
significantly longer, averaging from 8.5 to 12.5 d. These
early-born rod precursors seemed to initiate rhodopsin expression in a
manner that was not correlated temporally with the terminal mitosis.
For these cells, onset of rhodopsin expression
appeared approximately synchronous with the later-born cells,
suggesting a synchronous recruitment to the rod cell fate controlled by
environmental signals. To examine this possibility, we conducted
heterochronic cell mixing experiments and examined effects on the
kinetics of rhodopsin onset and induction to the rod cell fate in
early-born precursors exposed to a late retinal environment. In these
experiments, we demonstrate, similar to previous studies (Watanabe and
Raff, 1990), that the delay between the terminal mitosis and rhodopsin
expression in early-born rod precursors is not shortened by exposing
these early cells to a late environment using a reaggregate culture
system. Furthermore, in contrast to the conclusions drawn in the
previous report, we conclude here that the percentage of
early-born precursors adopting the rod cell fate is also unaltered when
these cells are transplanted to a late environment in vitro
(see below). These in vitro experiments thereby suggest that
the early-phase rod precursors are intrinsically distinct from the
late-phase precursors with respect to both the kinetics of rhodopsin
synthesis and the ability to produce rod photoreceptors.
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Figure 7.
Two distinct phases of rod development in
vivo. Rod genesis is divided into two phases in
vivo: an early phase and a late phase. Depicted schematically
are the time lags from the terminal mitosis of rod precursors to the
50% final rhodopsin level for cohorts born from E15 through P5. For a
majority of rod precursors, those born after E19, the timing of
rhodopsin onset is tightly linked to the timing of cell cycle
withdrawal. For these cohorts, the steps leading from the terminal
mitosis to rhodopsin expression take an average 5.5-6.5 d. These cells
constitute the late phase of rod genesis. During the early phase of rod
genesis, those cells born before E19 appear to display lags
significantly longer than 6.5 d, and commence rhodopsin expression
approximately synchronously with the late phase cells in the early
postnatal period.
|
|
Long latency and temporal correlation between rhodopsin onset and
cell cycle withdrawal for the majority of rods in vivo
A long delay between photoreceptor birthdays and opsin expression
appears well conserved across many species (for review, see Cepko,
1996). Previous studies in mouse and rat have measured the time between
BrdU labeling and the first BrdU+,
rhodopsin+ cells to be 48-54 hr in neonatal rod
precursors (Watanabe and Raff, 1990; Liou et al., 1994). It should be
noted that the data presented here do not differ from these previous
observations. For example, as shown in Figure 4A, the
first P0-born rod precursors express rhodopsin between 2 and 3 d
after their terminal S-phase. The measurement of 5.5-6.5 d represents
instead the average, or median lag, for these late born
rods. Because 80% of such rods begin to express rhodopsin between 4 and 7 d after the terminal mitosis, we believe that the average
lag is a more informative measurement for studying these rods than is
the lag to the first rhodopsin-expressing cells. Furthermore, the
present report is the first demonstration of the strong temporal
correlation in rat between the terminal mitosis and rhodopsin onset for
the majority of rod precursors in rodent retinal development.
Distinct kinetics of rhodopsin onset for early- and late-born
rods in vivo
The early-born rods display apparently distinct kinetics of
rhodopsin onset relative to the terminal mitosis. For cohorts born on
E15 and E17, rhodopsin onset becomes measurable only after a lag of
significantly longer than 6.5 d (Table 4). The onset of rhodopsin
in cohorts born before E19 appears roughly synchronous with the onset
of rhodopsin expression in later-born cohorts. These data suggest that
cohorts born earlier than E19 wait longer than 6.5 d to initiate
opsin expression and that this onset is roughly synchronous with
later-born cohorts. However, an alternative interpretation involving
differential cell death should be considered. There is a formal
possibility that the E15 and E17 cohorts may follow similar kinetics of
rhodopsin onset relative to their terminal mitosis as is followed by
the postnatal cohorts (i.e., a 5.5-6.5 d lag). These kinetics may be
masked to our measurements until a massive cell death of nonrods occurs
in the postnatal period. However, >90% differential cell death would
be required to shorten the observed long lags to 5.5-6.5 d. At this
time, existing data argues against this interpretation, because all
approximations of differential cell death in the early postnatal period
are estimated to be far lower than 90% (Young, 1984; Voyvodic et al.,
1995; Alexiades and Cepko, 1997). For example, 20% amacrine cell death was found during the P2-P11 interval, and amacrine cells constitute a
significant fraction of nonrods born on E15 and E17 (Alexiades and
Cepko, 1997). As well, cell loss between P1 and P15 for the E17 cohort
was found to be ~22% (Fig. 2A, see supporting
table). On the basis of these measurements, therefore, the longer lag seen for the early embryonically born cohorts is unlikely to be attributable to cell death alone.
Early and late precursors appear intrinsically distinct
in vitro
Early-born precursors and late-born precursors differ with respect
to the frequency that these cells choose the rod cell fate in
vivo. For example, ~5% of surviving cells born on E14
differentiate as rods, as compared with ~80% on P2 (Fig.
3A). As well, as discussed above, the early-born precursors
demonstrate a significantly longer lag to rhodopsin expression relative
to the later-born cells in vivo. In a study by Watanabe and
Raff (1990), the onset of rhodopsin expression in a population of cells
that were labeled with BrdU at E15 was similarly found to be delayed
relative to a population that was labeled on P1 in vitro.
These investigators tested whether the long lag observed for E15
labeled cells was extrinsically or intrinsically programmed by
conducting heterochronic cell-mixing experiments in vitro.
They argued that the longer delay seen for E15 cells was intrinsically
programmed because mixing E15 cells with excess P1 cells did not speed
the onset of rhodopsin in the E15 cells. The results presented here
confirm these original observations and are consistent with a
cell-autonomous inhibitor of rod differentiation acting in these
cells.
In contrast to the conclusions drawn here, Watanabe and Raff (1990)
concluded that exposing E15 cells to the P1 environment resulted in a
40-fold increase in E15 precursors adopting the rod cell fate. The
basis for this conclusion was an increase from 0.008 ± 0.002 to
0.44 ± 0.05% of E15 cells that were rhodopsin-positive after
6 d in vitro with an excess of P1 cells (Watanabe and
Raff, 1990). Despite their conclusion, because their cultures were not followed to the completion of rhodopsin expression, one is unable to
distinguish between an acceleration in rhodopsin kinetics in a small
subset of E15 cells at 6 d in vitro and an overall
increase in recruitment to the rod cell fate. In the current study, the cultures were followed to plateau levels of rhodopsin expression in vitro, and no alteration in the percentage of early-born
rod precursors expressing rhodopsin was observed. We therefore conclude that the vast majority of early-born precursors are not influenced with
respect to commitment to the rod cell fate when exposed to a late
environment in vitro.
The possibility that E16-born cells continue to be influenced by
signals derived from other E16 cells in the heterochronic pellets
cannot be formally excluded. However, when examined after 5 d
in vitro, the E16 cells remained well dispersed and were
surrounded by a majority of P0-derived cells (Fig.
6D) (Watanabe and Raff, 1990). These observations
lead us to favor the interpretation that the environment seen by the
E16-born cells and that of the P0-born control cells are highly
similar. Despite being placed in similar environments, these two cell
populations display distinct behaviors with respect to both rod genesis
and the kinetics of rhodopsin expression as discussed above. These
observations therefore suggest that the E16-born precursors and the
P0-born precursors are intrinsically distinct.
Characterizing the steps in rod development
For the majority of rod precursors, the onset of rhodopsin
expression is temporally correlated with cell cycle withdrawal, yet a
long latency, 5.5-6.5 d, is required to execute the steps leading from
a multipotent progenitor to a rhodopsin-expressing cell. What are the
steps leading from cell cycle arrest to rhodopsin expression that
require 5.5-6.5 d? There are presently several candidate genes that
may be involved in some of these steps. Among these are Notch and
HES-1, both of which have been shown to be inhibitors of neuronal
differentiation including rod differentiation (Dorsky et al., 1995;
Tomita et al., 1996; Bao and Cepko, 1997). As well, Pax-6 and Chx-10
are both expressed in retinal progenitor cells and ultimately become
restricted to nonphotoreceptor cell types late in development (Liu et
al., 1994; Hitchcock et al., 1996). These genes, therefore, are also
good candidates as inhibitors of rod differentiation. Finally, a
recently discovered paired-type homeobox gene, Crx, is
photoreceptor-specific and activates transcription from regulatory
elements of photoreceptor-specific genes in vitro (Chen et
al., 1997; Furukawa et al., 1997). Crx is, therefore, a candidate
promoter of rod differentiation. The data presented in the current
report now provide a framework in which to order the activity of these
important molecules and others in the pathway that leads a postmitotic
retinal precursor to a committed rhodopsin-expressing cell.
| |
FOOTNOTES |
Received Dec. 12, 1997; revised March 4, 1998; accepted March 5, 1998.
This work was supported by National Institutes of Health Grants EY09676
and EY08064. We thank Dr. R. S. Molday for an ample supply of the
Rho4D2 antibody, and M. Zeidler for help with confocal microscopy.
E.M.M. also thanks in particular Diala Ezzeddine for helpful
discussions during the course of the work and for a thorough and
critical reading of this manuscript. As well, we thank Z.-Z. Bao, A. Chen, and D. Schulte for feedback on this manuscript, and members of
the Cepko/Tabin Lab for help while these experiments were
conducted.
Correspondence should be addressed to Constance L. Cepko, Department of
Genetics and Howard Hughes Medical Institute, Harvard Medical School,
200 Longwood Avenue, Boston, MA 02115.
| |
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J. Sen, S. Harpavat, M. A. Peters, and C. L. Cepko
Retinoic acid regulates the expression of dorsoventral topographic guidance molecules in the chick retina
Development,
December 1, 2005;
132(23):
5147 - 5159.
[Abstract]
[Full Text]
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D. R. Graham, P. A. Overbeek, and J. D. Ash
Leukemia Inhibitory Factor Blocks Expression of Crx and Nrl Transcription Factors to Inhibit Photoreceptor Differentiation
Invest. Ophthalmol. Vis. Sci.,
July 1, 2005;
46(7):
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[Abstract]
[Full Text]
[PDF]
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Y. Tabata, Y. Ouchi, H. Kamiya, T. Manabe, K.-i. Arai, and S. Watanabe
Specification of the Retinal Fate of Mouse Embryonic Stem Cells by Ectopic Expression of Rx/rax, a Homeobox Gene
Mol. Cell. Biol.,
May 15, 2004;
24(10):
4513 - 4521.
[Abstract]
[Full Text]
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T. Matsuda and C. L. Cepko
Inaugural Article: Electroporation and RNA interference in the rodent retina in vivo and in vitro
PNAS,
January 6, 2004;
101(1):
16 - 22.
[Abstract]
[Full Text]
[PDF]
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J. James, A. V. Das, S. Bhattacharya, D. M. Chacko, X. Zhao, and I. Ahmad
In Vitro Generation of Early-Born Neurons from Late Retinal Progenitors
J. Neurosci.,
September 10, 2003;
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[Abstract]
[Full Text]
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T. Inoue, M. Hojo, Y. Bessho, Y. Tano, J. E. Lee, and R. Kageyama
Math3 and NeuroD regulate amacrine cell fate specification in the retina
Development,
March 4, 2003;
129(4):
831 - 842.
[Abstract]
[Full Text]
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S. Schulz-Key, H.-D. Hofmann, C. Beisenherz-Huss, C. Barbisch, and M. Kirsch
Ciliary Neurotrophic Factor as a Transient Negative Regulator of Rod Development in Rat Retina
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September 1, 2002;
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[Abstract]
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N. L. Brown, S. Patel, J. Brzezinski, and T. Glaser
Math5 is required for retinal ganglion cell and optic nerve formation
Development,
July 1, 2001;
128(13):
2497 - 2508.
[Abstract]
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M. A. Dyer and C. L. Cepko
p27Kip1 and p57Kip2 Regulate Proliferation in Distinct Retinal Progenitor Cell Populations
J. Neurosci.,
June 15, 2001;
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[Abstract]
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J Hatakeyama, K Tomita, T Inoue, and R Kageyama
Roles of homeobox and bHLH genes in specification of a retinal cell type
Development,
January 4, 2001;
128(8):
1313 - 1322.
[Abstract]
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X. Zhang and X. Yang
Regulation of retinal ganglion cell production by Sonic hedgehog
Development,
January 3, 2001;
128(6):
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[Abstract]
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P. A. Yourey, S. Gohari, J. L. Su, and R. F. Alderson
Vascular Endothelial Cell Growth Factors Promote the In Vitro Development of Rat Photoreceptor Cells
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E. S. Green, M. D. Menz, M. M. LaVail, and J. G. Flannery
Characterization of Rhodopsin Mis-sorting and Constitutive Activation in a Transgenic Rat Model of Retinitis Pigmentosa
Invest. Ophthalmol. Vis. Sci.,
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M. J. Belliveau, T. L. Young, and C. L. Cepko
Late Retinal Progenitor Cells Show Intrinsic Limitations in the Production of Cell Types and the Kinetics of Opsin Synthesis
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M. Dyer and C. Cepko
p57(Kip2) regulates progenitor cell proliferation and amacrine interneuron development in the mouse retina
Development,
January 8, 2000;
127(16):
3593 - 3605.
[Abstract]
[PDF]
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M Hojo, T Ohtsuka, N Hashimoto, G Gradwohl, F Guillemot, and R Kageyama
Glial cell fate specification modulated by the bHLH gene Hes5 in mouse retina
Development,
January 6, 2000;
127(12):
2515 - 2522.
[Abstract]
[PDF]
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M. Belliveau and C. Cepko
Extrinsic and intrinsic factors control the genesis of amacrine and cone cells in the rat retina
Development,
January 2, 1999;
126(3):
555 - 566.
[Abstract]
[PDF]
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E. Morrow, T Furukawa, J. Lee, and C. Cepko
NeuroD regulates multiple functions in the developing neural retina in rodent
Development,
January 1, 1999;
126(1):
23 - 36.
[Abstract]
[PDF]
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P. K. Swain, D. Hicks, A. J. Mears, I. J. Apel, J. E. Smith, S. K. John, A. Hendrickson, A. H. Milam, and A. Swaroop
Multiple Phosphorylated Isoforms of NRL Are Expressed in Rod Photoreceptors
J. Biol. Chem.,
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Top
Abstract
Introduction
