Current Biology 23, 443–452, March 18, 2013 ª2013 Elsevier Ltd All rights reserved
http://dx.doi.org/10.1016/j.cub.2013.02.007
Article
Rheotaxis Guides Mammalian Sperm
Kiyoshi Miki1 and David E. Clapham1,2,*
1Howard Hughes Medical Institute, Department of Cardiology,
Manton Center for Orphan Disease Research, Boston
Children’s Hospital, 320 Longwood Avenue, Enders 1309,
Boston, MA 02115, USA
2Department of Neurobiology, Harvard Medical School,
Boston, MA 02115, USA
Summary
Background: In sea urchins, spermatozoan motility is altered
by chemotactic peptides, giving rise to the assumption that
mammalian eggs also emit chemotactic agents that guide
spermatozoa through the female reproductive tract to the
mature oocyte. Mammalian spermatozoa indeed undergo
complex adaptations within the female (the process of capacitation) that are initiated by agents ranging from pH to progesterone, but these factors are not necessarily taxic. Currently,
chemotaxis, thermotaxis, and rheotaxis have not been definitively established in mammals.
Results: Here, we show that positive rheotaxis, the ability of
organisms to orient and swim against the flow of surrounding
fluid, is a major taxic factor for mouse and human sperm. This
flow is generated within 4 hr of sexual stimulation and coitus in
female mice; prolactin-triggered oviductal fluid secretion
clears the oviduct of debris, lowers viscosity, and generates
the stream that guides sperm migration in the oviduct. Rheotaxic movement is demonstrated in capacitated and uncapacitated spermatozoa in low- and high-viscosity media. Finally,
we show that a unique sperm motion, which we quantify using
the sperm head’s rolling rate, reflects sperm rotation that
generates essential force for positioning the sperm in the
stream. Rotation requires CatSper channels, presumably by
enabling Ca2+ influx.
Conclusions: We propose that rheotaxis is a major determinant of sperm guidance over long distances in the mammalian
female reproductive tract. Coitus induces fluid flow to guide
sperm in the oviduct. Sperm rheotaxis requires rotational
motion during CatSper channel-dependent hyperactivated
motility.
Introduction
Marine invertebrate spermatozoa, such as those of sea urchin,
are guided by chemotaxis; egg-secreted peptides induce
changes in the arc of spermatozoan swimming paths that
increase fertilization success [1]. In mammals, the route to
the egg is more restricted, but it is widely assumed that follicular fluid contains chemoattractants [2]. Prompted by reports
of the presence of a large number of putative odorant G
protein-coupled receptors in spermatozoa, the synthetic odorants bourgeonal and lyral were proposed to mimic unknown
native chemoattractants of human or mouse sperm, respectively [3, 4]. However, because muscle contraction and ciliary
currents would disrupt gradients over long distances, chemotactic guidance would likely be relevant only as sperm
*Correspondence: [email protected]
approach the egg [5, 6]. Another hypothesis, thermotaxy, is
based on ovulation-dependent small temperature gradients
between the isthmus and ampulla [7].
Rheotaxis, observed in many aquatic species, is the orientation of cells and organisms within a gradient of fluid flow. Positive rheotaxis, the tendency of a cell to swim against the flow,
was first reported for spermatozoa over 100 years ago [8].
While ciliary current toward the distal oviduct was observed
in turtle, pigeon, and rabbit, central lumen counterflow prevented a consensus about the physiological relevance of rheotaxis, and it was concluded to be a laboratory artifact [9]. In
addition to cilia-directed flow, muscle contraction, tortuous
geometry, the propensity of sperm to stick to surfaces, and
high-viscosity fluids were recognized as factors complicating
sperm transport. Rothschild [10] noted that the sperm head
is attracted to surfaces (the wall effect), and suggested that
the ‘‘lighter’’ tail is subjected to more force in velocity gradients, resulting in the tail being dragged downstream with the
head pointed upstream [11]. More recent considerations
explain the wall effect as a consequence of hydrodynamics
in tubes [12].
Fluid viscosity in the female reproductive tract varies, with
the highest being in the cervical mucus (200 to 680 cP in humans [13]) and the oviductal isthmus [14]. Oviductal viscosity
varies during the estrus cycle. Around ovulation, ciliated cells
in the isthmus epithelium are covered by a dense, tenacious
mucus that disappears after ovulation [14]; estrogen-regulated
secretion of oviductal fluid increases 2- to 3-fold at estrus [15].
Fluid viscosity also affects sperm movement—as viscosity
increases, sperm flagella propagation velocity, wave amplitude, and energy expenditure decrease [16]. Sperm angular
speed (rotation around the long axis of sperm) also decreases
at higher viscosities, resulting in more planar movement [17].
In the complex process of capacitation (that is, the changes
that sperm undergo in the female reproductive tract that
enable them to fertilize the egg), sperm develop hyperactivated motility. Hyperactivation is characterized by higheramplitude, asymmetric waveforms of reduced beat frequency
[18], which enables high flagellar swimming speeds in viscous
solutions [19].
We have reinvestigated sperm taxis in vitro and in vivo using
new tools that enable control of hyperactivated motility and
enable the visualization of sperm movement in vivo. Here, we
show that sexual intercourse in mice induces robust secretion
and fluid flow within the oviduct. Sperm are directed by,
and swim against, this flow (positive rheotaxis). Importantly,
axial rotation requires CatSper channels for reorientation of
sperm in tangential flow. We conclude that rheotaxis is a major
influence in the long distance guidance of mammalian sperm
to the egg.
Results
Coitus Induces Fluid Flow from Oviduct to Uterus
Oviductal epithelia actively secrete fluid into the oviductal
lumen, as first reported in rabbits during estrus [20]. Uterine
fluid is most abundant (91 6 26 ml) on the day of coitus [21];
however the origin of fluid and the mechanism of its production
has not been well documented. We analyzed the accumulation
of oviductal fluid in the uterus in mice. Although the uterus
Current Biology, Volume 23
Supplemental Information
Rheotaxis Guides Mammalian Sperm
Kiyoshi Miki and David E. Clapham
Supplemental Inventory
1. Supplemental Figures and Tables
Figure S1, Related to Figure 1
Table S1, Related to Figure 4
2. Supplemental Movie Legends
3. Supplemental References
Figure S1, Related to Figure
1. Sperm Movement in the
Ex Vivo Oviduct
(A) Cell debris/globular cells
discharged from the oviduct
via the isthmus (UTJ). The
oviduct of a mated female
mouse was removed and
incubated in HTF medium for
5h. Scale bar: 200µm.
(B) Oviductal ligation near the
utero-tubal junction (UTJ).
Oviducts were dissected from
mice 60-90 min after mating;
one of the two oviducts was
ligated near the UTJ in order to
block fluid flow from the
ampulla (white arrows) to the
UTJ (red arrows). Scale bar:
1mm.
(C) Transgenic sperm in the
ampulla. DsRed2 (red, tail)
and EGFP (green, head)expressing sperm in the
ampulla; unligated oviduct
under fluorescence
microscopy. This particular
spermatozoon did not undergo
the acrosomal reaction
(retained acrosomal EGFP).
Table S1, Related to Figure 4. Viscosity of Luminal Fluid and Media
Source
Rabbit, Oviduct [1]
Oviduct [2]
Mouse, Uterus, 4 h after coitus
Mouse, Uterus, 8 h after coitus
Mouse, Uterus, 4 h after coitus
Mouse, Uterus, 8 h after coitus
HTF medium
0.2% Methylcellulose in HTF
0.3% Methylcellulose in HTF
0.5% Methylcellulose in HTF
a
Fraction
1,500 g x 5 min supernatant
?
Total fluid
Total fluid
18,000 x g, 5 min supernatant
18,000 x g, 5 min supernatant
e
n.a.
e
n.a.
e
n.a.
e
n.a.
Viscosity was greater than that of 99% glycerol at 22ºC.
Mean ± s.d. (n = 4); Only samples >100 µl were measured.
c
Not determined due to low volume (see Figure 1c-ii).
d
Mean ± s.d. (n = 3); Only samples >100 µl were measured.
e
Not applicable.
b
Viscosity (cP)
1.1
1.8
a
n.d.
b
81 ± 73
c
n.d.
d
2.4 ± 0.9
0.98
3.7
6.7
29.8
Supplemental Movie Legends
Movie S1. Fluid Flow in the Oviduct
(A) Fluid flow in the oviductal isthmus. An oviduct removed from a mated female mouse
was incubated at 37ºC. Movement of cell debris/globular cells in the isthmus was
observed through its transparent wall and recorded by a CCD camera at 30 fps. Left
and right of the field are oriented to uterus or ovary, respectively.
(B) Ciliary current in oviductal ampulla. An oviduct dissected from a mated female
mouse was mounted on a glass slide. Ciliary current in the ampulla was observed
through the transparent wall and recorded by a CCD camera at 60fps. Bottom right and
upper left of the field are oriented to uterus and ovary, respectively. Objective: 20x.
Movie S2. Sperm Rheotaxis In Vitro
(A) Uncapacitated mouse sperm left in the 'no-flow' condition.
(B) Uncapacitated mouse sperm in flow generated by convection (10 µm/s).
(C) Uncapacitated mouse sperm under uniform laminar flow (generated by a pipette; 12
µm/s). Sperm are incubated in M2 at 37ºC. 10x objective. 60 fps.
Movie S3. Mouse Sperm Rheotaxis In Vitro
(A) Uncapacitated mouse sperm in the 'no-flow' condition in M2 medium at 37ºC; 60
fps.
(B) Uncapacitated mouse sperm under fluid flow (16 µm/s) in M2 medium at 37ºC; 30
fps; 4x objective.
Movie S4. Human Sperm Rheotaxis In Vitro
(A) Uncapacitated human sperm in the 'no-flow' condition.
(B) Uncapacitated human sperm under fluid flow (10 µm/s). Sperm were incubated in
HEPES-HTF at 37ºC; 10x objective; 60 fps.
Movie S5. Sperm Movement in a Capillary Tube during Low Viscosity Fluid Flow
(A) Loading of mouse capacitated sperm into a glass capillary tube at 210 µm/s in
HEPES-HTF at 22ºC. 10x objective.
(B) Immotile, aggregated, and sluggish sperm are swept out of the capillary tube by
outward-directed flow at 68 µm/s at 37ºC; 20x objective; 30 fps.
Movie S6. Sperm Selection in a Capillary Tube during Fluid Flow
Capacitated mouse sperm were loaded as in Movie S5A; (A) immediately after loading;
(B) followed by addition of sperm-free media (HEPES-HTF) to the left end of the tube.
By increasing the distance sperm had to swim, this procedure resulted in selection of
active sperm. Sperm take longer reach to the end of the capillary against flow, but
sluggish sperm are eliminated. 77µm/s; 37ºC; 30 fps.
Movie S7. Enhanced Progressive Motility in Viscous Medium; No Flow
(A) Uncapacitated mouse sperm in HEPES-HTF (~3 min).
(B) Capacitated mouse sperm in HEPES-HTF (2 h) at 37ºC; 10x objective; 60 fps. Note
the transition from planar circular motion before capacitation to more linear paths during
capacitation.
Movie S8. Sperm Movement in a Capillary Tube during Viscous Fluid Flow
(A) Uncapacitated mouse sperm swim along the inside surface of the capillary tube;M2
media (0.3% (w/v) MC) at 37ºC; 58 µm/s; 20x objective; 30 fps.
(B) Capacitated mouse sperm swim in the center of the capillary; HEPES-HTF (0.3%
MC) at 37ºC; 61 µm/s; 30 fps.
Movie S9. Sperm Turning in Flowing Solution
The path of the turning sperm is marked by the dotted line. Uncapacitated mouse sperm
in M2 medium at 37ºC; 11 µm/s; 10x objective; 60 fps.
Movie S10. Headless Sperm Rheotax
Flow rate = 6.2 µm/s in uncapacitated conditions in M2 medium; 10x objective; motion
slowed 10 fold to 6 fps.
Movie S11. Sperm Lacking CatSper Channels Have Low Motility, Swim in Circles,
and Do Not Rotate at 37oC
(A) No flow; (B) flow = 15 µm/s, 4x objective; (C) flow 15 µm/s, 20x objective.
Uncapacitated sperm (HEPES-HTF for 5 min only); 60 fps. Sperm dissected from the
cauda epididymis of CatSper1-/- mice.
Movie S12. Lack of Rheotaxis in Sea Urchin Sperm Regardless of Fluid Flow
(A) No flow.
(B) Laminar flow 17 µm/s; 22ºC. Motion slowed 2-fold to 60 fps.
Supplemental References
1.
Hamner, C.E., and Fox, S.B. (1968). Effect of oestrogen and progesterone on
physical properties of rabbit oviduct fluid. Journal of Reproduction and Fertility
16, 121-122.
2.
Menezo, Y., and Guerin, P. (1997). The mammalian oviduct: biochemistry and
physiology. European Journal of Obstetrics, Gynecology, and Reproductive
Biology 73, 99-104.
Current Biology Vol 23 No 6
444
A (i)
(iii)
(ii)
(iv)
C (i)
(ii)
(iii)
Recovered fluid (mg)
B
D
Hours after mating
p = 0.0007
G
p = 0.008
# sperm / ampulla
Recovered fluid (mg)
F
Flow speed (µm/s)
E
Vehicle BrCr
in Unligated Ligated
vivo
ex vivo
noticeably swells at estrus, no appreciable fluid was recovered
from ovulated females at any time point up to 24 hr after estrus
(Figure 1B). When females were mated with males, we
observed a slight swelling of the uterus 20 min after mating
and were able to collect 22 6 4.8 mg of fluid (primarily ejaculated semen; Figures 1Aii and 1Ci). Moreover, fluid in the uterus
increased over time to a peak of 87 6 13.9 mg at 8 hr (Figure 1B;
the rate of secretion appeared to be highest within the first
4 hr after mating). This increase in secretion above baseline
at estrus required coitus; unmated control mice did not exhibit
a significant increase in uterine fluid. Because in vitro culture
showed that the uterine accumulation included oviductreleased cell debris and globular cells (see Figure S1A
available online), we surmised that some portion of this fluid
in vivo originates from oviductal fluid.
Oviductal fluid pH remains fairly constant during the follicular phase of the estrus cycle. During ovulation in rhesus
monkeys, bicarbonate secretion into the oviduct increases
its pH from 7.1–7.3 to 7.5–7.8 [22]. Mouse uterine fluid recovered within 4 hr after coitus was a highly viscous colloid of
seminal fluid, cell debris, and globular cells, but the fluid’s
viscosity decreased over time (Table S1). Beginning a few
hours after coitus, significant secretion of bicarbonate-rich
oviductal fluid increased the pH to w8.3, reduced mucous
viscosity, and washed out viscous cellular debris (Figure 1C;
Movie S1A). Cilia sweep fluid and the egg toward the uterus,
Figure 1. Increased Fluid Production and Oviductal
Flow after Mating
(A) The mouse uterus swells after mating. No fluid
was recovered from the uterus at ovulation (i). Fluid
from uteri removed immediately after mating
increased by 37 mg, consistent with the weight
(volume) of sperm fluid (ii). Fluid volumes increased
markedly in uteri 8 or 12 hr after mating (120 or
100 mg, respectively) (iii and iv). Scale bar represents 1 cm.
(B) Accumulated uterine fluid was collected at the
times indicated after mating. Fluid weights (volumes)
from mated (red) and unmated (blue) females are
plotted.
(C) Uterine fluid was collected from females 20 min
(i), 4 hr (ii), or 8 hr (iii) after mating, and the fluids
were centrifuged.
(D) Fluid flow in the oviduct (ampulla) 4 hr after
mating. Migration of globular cells and debris was
traced over 3 s; the closed circle marks the endpoint.
The central diagonal structure is a mucosal fold in the
oviductal wall. Oviductal isthmus is seen at bottom
right. Scale bar represents 50 mm. See Movie S1.
(E) Average globular cell migration speed = 18.0 6
1.6 mm/s at 22 C.
(F) Bromocriptine administration before mating
inhibits fluid secretion measured 8 hr after mating.
(G) DsRed2-expressing sperm counts in the ampulla
in vitro and ex vivo. Ex vivo: a portion of ampulla was
dissected from females 4–5 hr after mating, and
DsRed2-expressing sperm were counted. Oviducts
were removed and ligated, or left unligated, near
the UTJ 60–90 min after mating and then incubated
at 37 C for 2–3 hr. The number of sperm reaching
the ampulla in oviducts without an ovary averaged
31 6 6. The increase in sperm number over the in vivo
situation may result from manipulation or a change in
hormone status perhaps detaching some sperm
from the isthmal crypts and facilitating sperm migration. All graphs: mean 6 SEM (black bars) and
median with interquartile ranges (green boxes).
reducing the attachment of cells to the wall of the oviduct
(Figure 1D; Movie S1B). Ampullar flow, measured 3–4 hr after
coitus, was 18.0 6 1.6 mm/s (Figure 1E). In summary, coitus
induces a dramatic increase in oviductal fluid secretion that
lowers viscosity and clears debris from the oviduct. We then
asked what factors might mediate this increased oviductal
secretion and flow.
Genital stimulation and coitus evoke complex neurological
responses. To simplify our task of determining factors that
might increase oviductal flow, we considered known secreted
hormones that were (1) elevated in serum after sexual stimulation or coitus, (2) known to effect fluid secretion across
epithelia surfaces, or (3) known to alter fertilization success
in mice in which the gene had been deleted. These criteria
narrowed our candidate hormones to prolactin, oxytocin,
and vasopressin. Prolactin is released from the anterior pituitary in response to sexual arousal circuitry that releases dopamine onto neuroendocrine cells. Prolactin, in turn, counters
dopamine’s mediation of sexual arousal and reduces sexual
receptivity [23]. In humans, plasma prolactin concentrations
are increased for >1 hr following orgasm induced by masturbation or sexual intercourse [24]. Female mice lacking the
prolactin receptor gene have multiple reproductive defects,
including reduced fertilization rate [25]. Finally, prolactin also
has a plethora of effects on secretory epithelia [26]. To test
whether prolactin could mediate oviductal secretion, we
Sperm Rheotaxis
445
A
33.5䉝 (surface)
37
Figure 2. Sperm Rheotaxis In Vitro
H
(A) Diagram of convective flow pattern observed
in warmed media.
(B) Laminar flow (1.5 ml/hr) from a rectangular
capillary at 37 C.
(C–F) Trajectories of mouse (in C, 3 s; in D, 4 s),
and human (in E and F, 5 s) sperm in flow,
analyzed by CASA. Scale bars represent 200 mm
(C and D) and 100 mm (E and F).
(G) Positive rheotaxis is defined here as sperm
movement against fluid flow within 622.5 of
the forward vector of sperm movement.
(H) Rates of rheotaxis for mouse and human
sperm (mean 6 SD) in five and three independent
experiments, respectively. Total sperm count is
shown in parentheses. See Movies S3 and S4.
No flow
D
p = 10-6
n=5
(288)
flow
p = 0.0002
n=3
(270)
E
Human
G
(bottom)
Mouse
C
B
No flow
F
n=5
(467)
n=3
(240)
flow
and no fertility phenotypes in TRPC5, V1,
V3, A1, or M8 knockout mice [29–33].
Furthermore, we failed to detect any of
these TRP currents in voltage-clamped
mouse spermatozoa. Most tellingly,
we noticed that temperature gradients create convection
currents that influence the direction in which sperm swim. As
the chamber warms the medium from the bottom surface, fluid
flows in well-recognized convection patterns, reaching
a steady state in which fluid cycles from bottom to top and
then down the sides of the chamber (Figure 2A). Sperm consistently swam against this flow (positive rheotaxis), whether
along the top liquid surface (in the direction of increasing
temperature) or bottom (in the direction of decreasing temperature) of the dish (Movies S2A and S2B). Solutions flowing
through a pipette into a dish at constant temperature initiated
identical positive rheotaxis (Movie S2C). To quantify rheotaxis,
we defined rheotactic movement as movement within 622.5
of the forward vector of sperm movement (swimming directly
upstream; small particles defined the local vector of flow; Figure 2G). Providing the baseline in static media, we observe
12.5% of sperm in the field swim in the direction specified by
the 45 arc centered along the sperm longitudinal axis. Without
flow, mouse and human sperm swam in random directions
(Figure 2H).
Mouse
administered the dopaminergic receptor agonist bromocriptine (BrCr) 11 hr before coitus to inhibit prolactin secretion
from the anterior pituitary. BrCr treatment reduced mouse
uterine fluid accumulation (Figure 1F). In contrast, administration of the VR2 receptor antagonists oxytocin (atosiban) or
vasopressin (tolvaptan) did not suppress oviductal secretion.
We conclude that prolactin initiates the increased oviductal
fluid flow to the uterus after coition. We next asked whether
oviductal flow affected sperm movement.
Sperm Migration in the Oviduct
Ejaculated sperm migrate through viscous fluid to bind to
epithelial cilia in the oviductal isthmus [6]. We tested whether
sperm then detached from the cilia and swim to the ampulla
after the mucus fluid had been cleared. Taking advantage of
genetically modified sperm, in which enhanced green fluorescent protein (EGFP) is expressed in the sperm acrosome and
DsRed2 fluorescent protein is expressed in sperm mitochondria [27] (Figure S1C), we found that 9 6 1 sperm reached
the ampulla 4–5 hr after mating (Figure 1G; in vivo). In order
to test whether rheotaxis was functional ex vivo, oviducts
were dissected from mice 60–90 min after mating. By this
time, sperm had reached the isthmus, and in the next 2–3 hr,
they migrated from the isthmus to the ampulla. Importantly,
sperm in the isthmus migrated successfully to the ampulla
ex vivo in the absence of an ovary, a source of potential chemoattractants (Figure 1G). However, sperm transport was
limited when oviducts were ligated near the uterotubal junction
(UTJ) to block fluid flow (Figure S1B), as assessed by observation of DsRed2-tagged sperm upstream of the ligation site and
by the lack of discharge of cell debris and globular cells. Sperm
motility in the blocked oviduct appeared normal, but there
were w3-fold fewer sperm in the ampulla of the ligated
oviducts (11 6 2), indicating that fluid flow is a factor in optimal
sperm transport ex vivo.
Spermatozoan Rheotaxis
The suggestion by several groups that transient receptor
potential (TRP) channels are present in spermatozoa and
might mediate sperm thermotaxis [28] motivated our initial
studies to examine sperm movement in a thermal gradient.
However, we found no effects on sperm motility or fertilization
Human
Rheotaxis of Uncapacitated and Capacitated Sperm in
Low-Viscosity Media
Because sperm capacitation can vary with respect to position
and time in the female reproductive tract [18], we evaluated
sperm rheotaxis in both uncapacitated and capacitated
mouse sperm. Ejaculated sperm swim to the oviductal isthmus
through the UTJ within 1–2 hr, but most require several more
hours to reach the ampulla [6]. When uncapacitated mouse
or human sperm were added to the convective flow chamber,
68% and 51% of the actively motile sperm exhibited rheotaxis,
respectively (Figures 2C–2F and 2H; Movies S3 and S4). To remove confounding results due to potential thermotaxis, we
repeated these experiments in isothermal fields of flowing
media. Uncapacitated sperm were placed in shear flow (Reynolds number [Re] = 0.002) via a 1 3 0.05 mm (w 3 h) capillary
tube (Figure 2B). As expected, spermatozoa swam against the
direction of flow (positive rheotaxis; Movie S2C), whereas in
controls (no flow) sperm movement was randomly directed
(Movie S2A).
Laminar flow in a chamber varies due to surface drag;
velocity is highest in the center of the stream and falls to
Current Biology Vol 23 No 6
446
Figure 3. Rheotaxis of Mouse Sperm in Low- and
High-Viscosity Media
Outward flow
A
exiting
Loading
Inward flow
inner diameter: 80 µm
exiting
B
Uncap
Exiting
FlowOUT
a
Exiting
(814)
c
(509)
FlowIN
FlowOUT
High viscosity
(804)
(587)
Loading
Cap
C
Low viscosity
Loading
(633)
a
(531) c
(843)
(877)
b
d
(825)
FlowIN
(616)
(697)
b
(544)
Sperm (%)
Sperm (%)
Motile
Sluggish
Immotile
a minimum next to the walls [34]. The flow gradient was
quantified by measuring 1 mm spherical beads placed in the
fluid as markers. Velocity profiles in fluid chambers are parabolic and approach linearity only near a small portion of their
profile near a boundary. By tracking beads, we found that
flow speed increased in the range of 0–300 mm from the
surface. Bead velocity in the range of 0–80 mm could be
approximated as w1.45x, where x is the distance in microns
from the surface (least-squares r2 = 0.98). The swimming
tracks of added sperm appeared as ‘‘threads’’ (similar to those
in Figures 2D and 2F), swimming w9 mm from the surface (flow
velocity = 14.5 mm/s). In this low-viscosity fluid, capacitated
mouse sperm that swim into the higher velocity of the central
stream are swept downstream. Sperm swim near the surface
where flow rate is slow, and those that stray away from the
surface are swept away by the faster, more central flow under
these conditions.
The lumen of the mouse oviductal isthmus consists of
a narrow central channel surrounded by pockets formed by
deep transverse mucosal folds. Sperm in the mucosal folds
can access the central channel to migrate in the oviduct. To
simulate the narrow central channel, sperm were capacitated
for 2–3 hr in human tubal fluid (HTF) medium (cooled to 22 C
to limit sperm motility) and loaded into the capillary (Figure 3A;
Movie S5A). The capillary was then transferred to a 37 C
chamber where sperm resumed more active motility. Under
outward-directed flow (FlowOUT; w15 nl/min or w50 mm/s),
sperm motility was recorded and sperm that had been swept
out the end of the capillary counted. Whereas 44% of the
sperm loaded into the tube were immotile, 78% of sperm
swept from the tube were immotile (Figure 3B; Movie S5B).
Conversely, when sperm were slowly sucked into the
pipette (inward flow; FlowIN; w15 nl/min or w50 mm/s; 39%
motile), 93% of sperm were active and moving against fluid
flow, leaving immotile and some sluggish sperm behind (Figure 3B; Movie S6). Thus, gentle flow concentrates, or selects,
viable sperm. Uncapacitated sperm showed essentially the
same results (Figure 3B). In short, sperm swim upstream in
a capillary mimicking flow rates in the oviductal lumen, a direction that also moves the oocyte toward the incoming
d
(A) Capacitated (Cap) or uncapacitated (Uncap)
sperm were suspended in medium with (highviscosity) or without (low-viscosity) 0.3% (w/v)
methylcellulose. Sperm were loaded into a capillary (at 22 C), and positive (outward flow; w15 nl/
min) or negative (inward flow; w15 nl/min) pressure was applied (at 37 C). The most immotile
and sluggish sperm were present in the central
stream (w50 mm/s flow rate), whereas most of
the motile sperm swam near the wall where the
flow rate was lowest.
(B and C) Sperm were categorized as motile,
sluggish, or immotile. Sperm movement near
the end of the capillary was observed under
outward (FlowOUT) or inward (FlowIN) flow in the
low- (B) or high-viscosity (C) medium (mean 6
SD, n = 5; total sperm counts are shown in
parentheses). Statistical significance was compared between the number of motile sperm
in FlowOUT and FlowIN. p values of each experiment (compare error bars marked as a, b, c,
and d) were smaller than 1026. See Movies
S5, S6, and S8.
sperm and eventually to the uterus. Thus, oviductal secretion
and flow clear the oviduct and increase pH to induce hyperactivation; rheotaxis selects for motile sperm and ensures that
eggs are fertilized while still in the oviduct. As proposed by
Suarez and colleagues, slow release from reservoirs of quiescent sperm or sperm attached to epithelial or ciliary surfaces
may provide a steady supply of sperm as this process is
repeated over time [6].
Sperm Movement in High-Viscosity Media
During estrus, high-viscosity mucus is an obstacle to sperm
progress through the isthmus lumen. Assuming, as a first
approximation, that the fluid is Newtonian, we measured the
viscosity of secreted and accumulated lumenal fluid in the
uterus, in which oviductal mucus, cell debris, and ejaculated
semen were present. The viscosities of this fluid averaged
81 6 73 cP compared to the viscosity of clear fluid supernatants (2.4 6 0.9 cP; Table S1). One caveat however, is that
the fluid is thixotropic (viscosity decreases with shear) and
thus likely non-Newtonian. Nevertheless, we estimate that
the viscosity of lumenal fluid decreases to as low as 2–3 cP
upstream of the congestion of mucus and cell debris in the
isthmus. Therefore, we used media containing 0.3% methylcellulose (MC) (6.7 cP; Table S1) to mimic the environment of
the oviductal lumen. As seen in Figure 4A and Movie S7A, uncapacitated mouse sperm swim in circles in viscous medium
regardless of priming by bicarbonate ion in HTF medium
(w3 min), which increases flagellar beat frequency within
30 s [35]. The majority of uncapacitated sperm exhibit planar
swimming (Figure 4A), with one side of the head facing the
surface. This swimming behavior may increase the chance
for uncapacitated sperm to stick to oviductal epithelial cells
more frequently, a delay that might promote acquisition of
the capacitated state [36].
Sperm Rotate after Capacitation in Viscous Media
When sperm are observed under phase-contrast microscopy,
reflection intensity varies sinusoidally as the sperm head
rotates, with the highest intensity corresponding to the head
in the position vertical to the surface [13] (Figure 4C). We
Sperm Rheotaxis
447
Figure 4. Sperm Rotation during Capacitation
Brightness (a.u.)
D
A
250
150
uncapacitated
0
0.5
1.0
E
B
Brightness (a.u.)
Seconds
250
150
capacitated
0
0.5
(A and B) Sperm trajectories in viscous solution: sperm were incubated with 0.3% (w/v)
methylcellulose-containing HEPES-buffered HTF
medium for w3 min (A: uncapacitated) or 2 hr
(B: capacitated). Scale bar represents 200 mm.
See Movie S7.
(C) The head of the spermatozoa is dark on
phase-contrast images due to its relatively flat
structure (upper panels). Sperm head light reflectance increases on phase-contrast images when
the sperm heads orient vertically to the surface
(lower panels).
(D and E) Representative results of the sperm
head-rolling (rotation) analysis. Uncapacitated
(D) or capacitated (E) sperm were incubated in
HTF medium for 3 hr and analyzed (a.u. indicates
arbitrary units).
(F) Increase of rotation rate during capacitation
(6SD; w70 sperm for each time point).
1.0
Seconds
F
Rotation rate (Hz)
C
8
6
fertilization site faster than uncapacitated sperm in high-viscosity solution.
Increased Rotation in Turning
As noted above, sperm often swim close
4
to surfaces due to the hydrodynamic
wall effect [10]. We observed that sperm
eventually leave the surface, are
Vertical
2
exposed to faster flow and pushed
downstream, but then they return to
Incubation (min)
the surface and again orient against
fluid flow (Figure 5A; Movie S9). We
0
50
100
150
200
noticed rapid sperm rotation (3.8 6
0.2 Hz; 6SEM) when the sperm turns in
plotted sperm head reflection intensity as a function of time response to fluid flow at a position perpendicular to the direcand call it ‘‘rotation frequency,’’ because it reflects the spin- tion of fluid flow (11 out of 14 analyzed sperm; Figure 5B),
ning of the sperm head and tail around a longitudinal axis. followed by a return to normal rotation frequency (3.0 6
Rotation frequency was 1.7 6 0.1 (6SD)-fold higher in capaci- 0.2 Hz; 6SEM). Time-course analysis of turning sperm retated (Figures 4E and 4F) than uncapacitated (Figures 4D vealed a peak rotation rate of 7.1 Hz during turning, compared
and 4F) sperm. During capacitation, the linear progress of to 4.5 Hz prior to turning (Figure 5C, red). In contrast, the rotathe movement is increased by rotation (Figure 4B; Movie tion frequency of sperm in steady-state positive rheotaxic
S7B). In flowing high viscosity media, uncapacitated sperm movement was constant during steady flow and indistinswim in a corkscrew path, and the head does not rotate guishable from sperm in ‘‘no-flow’’ conditions (Figures 5C
when closest to the wall (Movie S8A). This motion may be and 5D). To further examine the propensity of sperm to turn
boundary-driven (thigmotaxic) rather than rheotaxic. Sperm into a stream we examined sperm at the confluence of two
rotation after capacitation may enhance detachment from flow streams 45 apart (Figure 5E). In these situations, sperm
surfaces. Once in the stream, linear progress is enhanced; rotation frequency significantly increased and resulted in
the head rotates and the sperm progression pattern oscillates sperm being directed into the stronger stream. Since inertia
around a motion-oriented parallel to the flow (Movie S8B). is negligible in the microscopic range [12] but governed by
Linear velocity (migration speed toward the capillary end) of overdamped, zero Reynolds number dynamics, the system
uncapacitated and capacitated sperm was 36 6 6 and 54 6 will be governed not by momentum, but rather by relative
8 mm/s (mean 6 SD, n = 11), respectively, under comparable forces. Our conclusion is that sperm increase their rotation
flow conditions in viscous media (Movie S8). In summary, rate when they encounter tangential (side) forces.
sperm rotation after capacitation enables sperm to swim
into the main fluid stream (Figure 3C; Movie S8B). At the Mechanisms: Tail Rotation Is Required for Rheotactic
boundary (chamber floor), the fluid velocity approaches zero, Movement
but as the sperm head moves up into the fluid, sperm experi- Orientation in a flowing stream has been described for
ence a difference in flow velocity that rotates the cell. This mammalian sperm [11, 35] and bacteria [37]. Some mechapreviously unappreciated capacitation-induced movement is nistic models of this orientation assume stronger hydrodyan important aspect of motion, in addition to the increased namic drag on the head (sperm) or cell body (bacteria) than
motility of sperm in viscous medium reported by Suarez the flagellum that orients the front of the cell toward the
et al. [19]. Thus, the gain in forward progression in the surface, resulting in the tail being dragged downstream with
oviductal lumen enables capacitated sperm to reach the the head and cell body pointed upstream [11, 37]. To separate
Flat
Current Biology Vol 23 No 6
448
Figure 5. Increased Sperm Rotation Rate in
Fluid Flow
A
Perpendicular
1s
1s
0.5 s
Rotation rate (Hz)
B
Rotation rate (Hz)
C
turning sperm
}
(A) Representative uncapacitated spermatozoan
turns in response to fluid flow. Images were recorded at 0.5 s intervals; black arrows mark
a turning sperm. Flow direction, in all panels, is
indicated by the green arrow. See Movie S9.
(B and C) Rotation rate of individual turning
sperm; average of two 1 s periods plotted at
0.5 s intervals (some points overlap and are
shown as single dots) (B) or sequentially over
time (C). Red line in (C) indicates a turning sperm;
other lines indicate sperm swimming in a straight
line against fluid flow.
(D and E) Conditions that change sperm rotation
rate. Uncapacitated sperm were incubated at
37 C, and the rate of rotation measured in no
flow, steady flow (14 mm/s), or convergent flow
(shown in E; 6SEM, n = 35).
non-turning sperm
fluid flow; their trajectories were similar
to those of mouse sperm lacking CatS2
Seconds
per channels or uncapacitated sperm in
D
p = 0.0003 E
high-viscosity medium (Figure 6B;
Movie S12A). Regardless of fluid flow,
convergent
p = 0.0004
these sea urchin sperm swam in circular
flow
planes (Movie S12B), consistent with
the behavior of sperm from marine
invertebrates that are not guided by
marine currents but swim in concentric
p = 0.0001
circles until encountering chemoattractants that turn and widen their arcs of
pipette circular motion [39].
Prior studies show the sperm
Turn Linear
flagellum oscillates in a single plane
convection
when tethered at its head on a glass
coverslip [35]. In order to better understand the motion of sperm in three
dimensions during free swimming, we
recorded a sperm flagellum every
50 ms and aligned these waveforms
by horizontal translocation of the
the differential influence of drag on head and tail, we examined swimming direction axis (Figure 7A). During free swimming,
swimming headless sperm, which represent a minute fraction flagella crossed the x axis at w20% of their entire length as
(<0.1%) of sperm from normal mice. Of these, 82% (14 out of measured from the head (Figure 7A, right). This point is the
17) swim against fluid flow (Movie S10), indicating that the minimum of flagellar excursion (Figure 7, arrowhead) and
head does not act as an aft rudder and is not required for moves along the x axis with minimal y deviation or yaw into
normal orientation—in other words, the rotating tail alone the plane (z axis). If sperm do not rotate, flagellar trajectories
trace a triangular plane, resulting in minimal rheotaxis.
can exhibit positive rheotaxis.
Sperm lacking any one of five CatSper subunits (CatSpers1–4, Sperm rotation produces trajectories that trace a symmetric
CatSperd) exhibit identical phenotypes [38], fail to develop cone-like path (Figure 7B), consistent with previous observafunctional CatSper Ca2+ currents, and lack hyperactivated tions of human spermatozoa that trace out a cone-shaped
motility. Uncapacitated sperm lacking ICatSper usually helical pattern and have an extended elliptical cross section
swam in a counterclockwise circular plane on the bottom when their heads are immobilized by attachment to a micropisurface, as observed from the top (Figure 6A; Movie S11A). pette, but allow the tail to exhibit flexible three-dimensional
This swimming behavior did not change regardless of fluid movement [40]. Tangential forces on the tail anterior to the
flow (Figure 6A; Movies S11B and S11C), indicating that the minimum of flagellar excursion will produce a clockwise force
CatSper channel was required for effective rheotactic (as seen from above), whereas those on the posterior tail
responses to fluid flow. Presumably, calcium influx through produce a counterclockwise force. The posterior tail receives
CatSper channels enhances sperm rotation, which changes stronger tangential force due to its higher proportion of the
circular planar movement to rotational swimming. Thus, rota- flagellar length (w80%, presenting a greater cross section to
tion is an essential sperm motion for proper rheotaxis, and the force of fluid flow), pointing the sperm upstream (FigCatSper channels contribute to this response. Interestingly, ure 7B). If the sperm were fixed at the point of minimal
S. droebachiensis sea urchin sperm did not swim against excursion, this would be analogous to a weather vane that is
Rotation rate (Hz)
1
Sperm Rheotaxis
449
A
No flow
A
CatSper KO
B
B
No flow
Sea Urchin
flow
Figure 6. Without Rotation, Mouse and Sea Urchin Sperm Swim in Circles
Trajectories of uncapacitated sperm from CatSper1 knockout mouse (A: 3 s)
or sea urchin sperm (B: 2 s). Fluid flow was generated by convection or via
pipette, and sperm trajectories were analyzed by CASA. Scale bar in (A)
represents 200 mm; scale bar in (B) represents 50 mm. Green arrows show
the direction of fluid flow. See Movies S11 and S12.
divided by an axis of rotation into a small anterior and a larger
posterior surface that receives stronger tangential force and
thus guides the weather vane into the wind. A freely motile
sperm in a homogeneous flow is not constrained to be
stationary at any point and therefore is not subject to torque,
but it is subjected to differences in flow velocity along its
length.
Discussion
We propose that rheotaxis, assisted by factors inducing hyperactivated motility (alkaline pH and, in humans, progesterone), is a major mechanism guiding sperm in mammalian
female reproductive tracts. This rheotaxis is not related to
sensory mechanisms and guidance responses as it is in large
animals such as fish but a consequence of rotational motion
potentiated by calcium entry via CatSper channels. Three
arguments support sperm rheotaxis as a major guidance and
selection mechanism. First, coitus triggers substantial fluid
secretion into the oviduct, thus increasing lumenal flow
directed by cilia and muscle contractions from ampulla to
uterus. Second, positive rheotaxis of mature spermatozoa
from mice and humans is observed in situ and in vitro. Finally,
capacitating conditions and physiological fluid flow overcome
the wall effect and sperm adhesion—these factors accelerate
sperm rotation that in high viscosity fluid evokes a change
from planar circular swimming to directional swimming. In
contrast, the most popular hypothesis for taxis, chemotaxis,
lacks both bona fide chemoattractant candidates and cognate
receptors in mammals. We cannot, however, exclude a role for
chemotaxis—if it occurs, it is most likely relevant over short
distances. Similarly, we conclude that our results cannot be
explained by thermotaxis. In the study of Bahat et al. [41],
only w5% of cells respond to temperature gradients, 8-fold
less than the percentage of sperm responding to rheotaxis in
our experimental system (w40% of the population; Figure 2).
Most importantly, we observed that sperm swam upstream
even though there was a descending temperature gradient.
Figure 7. Model for Sperm Rheotaxis
(A) The flagellum of an unattached sperm swimming against fluid flow was
traced every 50 ms and aligned by horizontal movement in the swimming
direction (dotted line). Arrowhead indicates point of minimal excursion of
flagellum (mef).
(B) Fluid flow (red arrows) reorients sperm (yellow arrows) into the flow to
reduce shear as the sperm rotate (orange arrow) and propel themselves
upstream. Rotation maps out a three-dimensional cone shape in space,
which orients sperm consistently into the flow (positive rheotaxis).
Again, this does not mean that true thermotaxis or thermal
effects on sperm motility are absent.
Fluid secretion in response to coitus has been relatively
unstudied. We found that there was little fluid secretion into
the oviduct and uterus at ovulation in mice. In contrast, coitus
induced a dramatic secretion of fluid that accumulated in the
uterus (pH w8.3), consistent with active extrusion of bicarbonate ion and water from the oviductal epithelium [42].
Interestingly, simulated coitus improves the success rate of
artificial insemination in mice [43]. In humans, sexual stimulation of females increases serum prolactin [24], and our initial
findings in mice suggest that this is mediated by bromocriptine-sensitive dopaminergic pathway control of the anterior
pituitary gland. Prolactin regulates water and electrolyte
balance in many organs (e.g., kidneys, fish gills [26]), perhaps
via activation or regulation of the cystic fibrosis transmembrane conductance regulator (CFTR) [23, 42] or aquaporins
[44]. With increased oviductal flow, globular cells from the
oviduct were also discharged into the uterus within 4 hr after
coitus. Apparently, ciliary motion and muscle contraction in
the isthmus pushes oviductal fluid to the uterus, in turn
washing away mucus and globular cells to clear the way for
sperm traffic. Sperm harbored in the lower isthmus then orient
and swim against this flow to reach the ampulla. Unlike
the isthmus, countercurrents in the center of the ampullar
lumen circulate fluid while ciliary currents push the fluid downstream into the oviduct [9]. Due to the hydrodynamic wall
effect [10], sperm moving locally along ciliated epithelium
may also swim against this ciliary current to reach the cumulus
oophorus complex.
Current Biology Vol 23 No 6
450
Sperm of marine invertebrates swim in a relatively unrestricted three-dimensional space, where the encounter of an
egg may be fortuitous. Chemotaxis substantially improves
the odds of reproduction of these species. In contrast, the
reproductive tract of mammals is restricted to a quasi-onedimensional tube with increasing pH (and progesterone in
humans) initiating hyperactivated motility to free sperm from
adherent surfaces and rheotaxis to guide the sperm toward
the oviduct. With up to 5 million mouse sperm deposited
in the females after ejaculation, only w20 sperm cells reach
the ampulla within several hours, similar to the success rate
of other mammals [40]. Oviductal flow has two advantages—it guides sperm to the egg but also selects for strong,
capacitated sperm. Mammalian sperm must penetrate
mucus, the viscoelastic matrix of the cumulus oophorus,
and the tough zona pellucida, requiring vigorous sperm for
successful fertilization. Also, directional flow sweeps the
egg down the oviduct, and sperm that swim against the flow
preferentially fertilize the egg while still in the oviduct. This
provides time for the egg to develop for implantation. From
this viewpoint, rheotaxis is a logical guidance and selection
mechanism.
Our results suggest that positive rheotaxis is due to the
spiral rotation of the sperm tail that orients sperm upstream,
and as such, it is decidedly an active process. Calcium entry
is required to trigger the increased amplitude of the distal
tail excursion that increases tangential force. At the same
time, increasing angular speed stabilizes the sperm’s swimming orientation due to increased motive force against the
surroundings. Because the hyperactivated flagellar waveform is both asymmetric and of larger amplitude [18], the
flagellum receives larger tangential forces, especially in
high-viscosity solution [6], which points the sperm into flowing solution. Of course, the underlying basis of motile force
and rotational motion is the basic adenosine triphosphate
(ATP)/dynein motor; calcium modulates this motor and other
components of the flagellar axoneme in ways that are not yet
understood.
In summary, we find that coitus induces oviductal flow
that clears the oviduct of cellular debris and provides rheotactic guidance for sperm. CatSper-mediated calcium entry
associated with capacitation and subsequent hyperactivated
motility potentiates and regulates the sperm rotation that is
crucial to guidance in the graded, complex fluid flow in the
oviduct.
Experimental Procedures
Mating and Collection of Uterine Fluid
All animal experiments were performed according to procedures approved
by the Institutional Animal Care and Use Committee of Boston Children’s
Hospital. Mice were housed in a 12 hr light (7 a.m.–7 p.m.)/dark cycle. Follicular growth and ovulation were stimulated in CD-1 female mice (Charles
River) at 8 to 9 weeks of age by intraperitoneal (i.p.) injection of pregnant
mare serum gonadotropin (5 IU, PMSG; Calbiochem) at 6:30 p.m., followed
48 hr later by i.p. injection of human chorionic gonadotropin (5 IU, hCG;
Calbiochem). In some experiments, BrCr (10 mg/kg, B2134; Sigma) was
subcutaneously injected 11 hr before mating. The mating period for
B6D2F1 males was set between 6:20 and 7:00 a.m. the morning after hCG
injection; the end of the mating period was t = 0. In some experiments,
atosiban (4 mg/kg) or tolvaptan (4 mg/kg) was injected i.p. just after mating.
The uterine horn was tied and dissected out with the oviduct and ovary, and
uterine fluid was collected by gently squeezing the cut end of the uterus with
forceps. The uterine fluid was the weight difference before and after fluid
collection. VR2 receptor antagonists for oxytocin (atosiban) or vasopressin
(tolvaptan) yielded 143 (659, SEM, n = 4) and 85 (616, SEM, n = 6) mg,
respectively, of uterine fluid 8 hr after mating.
Observation of Oviductal Fluid Flow
Oviducts were removed 3–4 hr after mating and placed in a 37 C chamber
containing M2 medium (2 ml; Specialty Media, Millipore). A cover glass
was placed over the oviduct, and fluid flow was recorded through the
oviductal wall (20 3 objective; CCD at 60 frames/s; 22 C).
Ex Vivo Sperm Migration
DsRed2-EGFP transgenic males [28] (B6D2F1-Tg; CAG/su9-DsRed2, Acr3EGFP RBGS002Osb) enabled clear sperm visualization. Sixty to ninety
minutes after the end of the mating period, oviducts were removed and
ligated just after the UTJ (ovarian side). Oviducts were gently washed in
HTF medium (Specialty Media, Millipore) to remove sperm attached to the
outside wall and then incubated (5% CO2) for 2–3 hr. A portion of the ampulla
was first cut with a fine scissors to prevent sperm from leaving the isthmus,
then the oviduct was stretched by trimming the mesosalpinx and placed on
a glass slide (14 each of ligated or unligated oviducts), and sperm inside
were counted under fluorescence microscopy. Samples in which there
were obvious sperm aggregates on either side of the ampulla were removed
from statistical analysis (two cases each; ligated and unligated oviducts).
Sperm Preparation
Mouse sperm were collected from the B6D2F1 male (Jackson Laboratories)
cauda epididymis and incubated for 5 min to allow sperm to swim out.
Centrifugation at 900 g (5 s) removed debris and large sperm aggregates.
Sperm were then incubated at a density of 2 3 106/ml in M2 media for 30 min
or HTF medium for 2–3 hr (5% CO2), yielding uncapacitated or capacitated
sperm, respectively. Cryopreserved human sperm were purchased from
Fairfax Cryobank. Actively motile human sperm were collected by the
swim-down procedure. Frozen sperm (w3 3 107) in egg yolks were
thawed and washed by suspension in 5 ml HTF, followed by centrifugation
at 500 g 3 5 min to obtain a sperm pellet. After two rinses, the pellets
were resuspended in 0.5 ml HTF. Half of the sperm suspension was overlaid
with a 0.5 ml Percoll cushion (80% HTF, 20% Percoll, 0.1 M NaCl).
Sperm were incubated for 45 min in a 37 C/CO2 incubator and sperm
swimming-down into a Percoll cushion were recovered by centrifugation
at 1,800 g 3 1 min. Sperm in the pellet were resuspended in 0.5 ml HTF
and incubated for 30–60 min.
Green sea urchin (Strongylocentrotus droebachiensis; ISF Trading,
Marine Biology Laboratory, Woods Hole, MA, USA) sperm were obtained
by intracoelomic injection of 0.5 M KCl. Eluted sperm were suspended in
artificial seawater (ASW) containing 486 mM NaCl, 30 mM MgSO4, 10 mM
MgCl2, 10 mM KCl, 10 mM CaCl2, 2.5 mM NaHCO3, and 10 mM HEPES
(pH 8.0).
In Vitro Sperm Rheotaxis
Sperm rheotactic movement was analyzed in convective and capillary flow.
For convective flow, 2 ml of culture medium and 1–2 3 105 mouse sperm or
2–4 3 105 human sperm were added to the 37 C chamber (Delta T culture
dish controller; Bioptechs). M2 or HEPES-HTF medium (92 mM NaCl,
2 mM CaCl2, 4.7 mM KCl, 0.2 mM MgCl2, 0.37 mM KH2PO4, 25 mM NaHCO3,
18.3 mM Na lactate, 2.78 mM glucose, 0.33 mM Na pyruvate, 0.4% [w/v]
bovine serum albumin [BSA], and 10 mM HEPES [pH 7.4]) was used for
mouse or human sperm, respectively. In some experiments, the medium
was supplemented with 0.3% (w/v) methylcellulose (MC) (M0512, 4,000 cP
in 2% solution; Sigma). Sperm swimming 3–5 mm from the rim were recorded by CCD camera (4 or 10 3 objective, mouse or human sperm,
respectively; KP-F31SCL; Hitachi) after a 10 min preincubation period that
allowed spontaneous dissociation of sperm clumps and stabilized convective flow. For control (no flow) and capillary flow experiments, 1 ml of
medium was overlaid by 1 ml of mineral oil and covered by a heated glass
lid (Bioptechs) to inhibit convection. A glass pipette of 1.2 mm (OD) rectangular glass capillary (w50 mm [h] 3 w1 mm [w]) was placed on the bottom
of the chamber, and the media was gravity fed at 1.5 ml/hr. Mouse sperm
(1–2 3 105) were transferred to the heating chamber and analyzed for
sperm rheotaxis at a constant 37 C. For sea urchin sperm, 2 ml of ASW
and 1–2 3 106 sperm were added to a 35 mm culture dish precoated with
0.1% gelatin to prevent sperm surface adhesion. ASW containing 1 mm
microbeads (Micromer; Micromod) was perfused at 3 ml/hr (22 C).
Measurement of Sperm Rheotaxis
Sperm motion in 3 s intervals was recorded by CCD at 60 frames/s,
processed using digital image processing software (XCAP; EPIX), and
analyzed using ImageJ. Swimming trajectories were via Computer Assisted
Sperm Analysis (CASA; http://rsbweb.nih.gov/ij/plugins/casa.html). Sperm
Sperm Rheotaxis
451
movement within 622.5 from the vector 2180 from the direction of fluid
flow was defined as rheotactic movement (Figure 2G). Sperm undergoing
rheotaxis were divided by the number of motile sperm; sluggish sperm,
swept out by convective flow, were excluded from the count.
Uncapacitated and Capacitated Sperm Movement in the Capillary Tube
Mouse sperm incubated in the HTF medium for 2–3 hr or M2 medium
for 30 min at 2 3 106/ml yielded capacitated or uncapacitated sperm,
respectively. Sperm were centrifuged at 900 g 3 1 min, resuspended in
HEPES-HTF or M2 medium with or without 0.3% (w/v) MC at 4 3 106/ml,
and loaded into the capillary by suction via an air-pressure microinjector
(IM-5B; Narishige; 22 C). Sperm in the capillary were transferred to the
37 C chamber containing 2 ml of HEPES-HTF or M2 medium with or without
0.3% MC at 37 C (w150 sperm for each experiment). In some experiments,
microbeads (1 mm) served as a flow indicator; a flow rate of w3 ml/mm2/min
provided w50 mm/sec flow in the cross-section imaged.
Viscosity Measurements
The viscosity of lumenal fluid and media was measured using the rolling-ball
method at 22 C. Variable glycerol concentrations were used as calibration
standards.
Statistical Analysis
Data analyses were performed using Microsoft Excel and GraphPad Prism.
Paired (Figure 5B) or two-tailed (all other figures) Student’s t test was used
to calculate statistical significance.
Supplemental Information
Supplemental Information includes one figure, one table, and 12 movies
and can be found with this article online at http://dx.doi.org/10.1016/
j.cub.2013.02.007.
Acknowledgments
We thank J.-J. Chung for help with CatSper12/2 mice, M. Delling and
S. Febvay for imaging assistance, Y. Miki for sample preparation, and the
Clapham laboratory for discussion. DsRed2-EGFP transgenic mice
(B6D2F1-Tg: CAG/su9-DsRed2, Acr3-EGFP: RBGS002Osb) were generously provided by M. Okabe, Osaka University, RIKEN BioResource Center.
This work was supported by NIH grant P30-HD18655 and the Bill and
Melinda Gates Foundation. We thank D. Babcock, University of Washington, and J. Kirkman-Brown and D. Smith, University of Birmingham, for
valuable comments on the manuscript.
Received: November 15, 2012
Revised: January 23, 2013
Accepted: February 4, 2013
Published: February 28, 2013
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