Rotation of endosomes demonstrates coordination of molecular motors during axonal transport

Abstract

Long-distance axonal transport is critical to the maintenance and function of neurons. Robust transport is ensured by the coordinated activities of multiple molecular motors acting in a team. Conventional live-cell imaging techniques used in axonal transport studies detect this activity by visualizing the translational dynamics of a cargo. However, translational measurements are insensitive to torques induced by motor activities. By using gold nanorods and multichannel polarization microscopy, we simultaneously measure the rotational and translational dynamics for thousands of axonally transported endosomes. We find that the rotational dynamics of an endosome provide complementary information regarding molecular motor activities to the conventionally tracked translational dynamics. Rotational dynamics correlate with translational dynamics, particularly in cases of increased rotation after switches between kinesin- and dynein-mediated transport. Furthermore, unambiguous measurement of nanorod angle shows that endosome-contained nanorods align with the orientation of microtubules, suggesting a direct mechanical linkage between the ligand-receptor complex and the microtubule motors.

Fig. 1. Transport by teams of motors results in rotational dynamics.

Schematic of a hypothetical endosome with three dyneins and one kinesin starts transporting in the configuration of the middle image (dashed box) and can transition to any of the states in conditions 1 to 6. In conditions 1 to 3, the translational motion is paused. In condition 1, the two previously bound dyneins unbind, and the cargo freely tumbles. However, the confinement of the axon restricts how far it will diffuse, making it difficult to detect significant diffusive translational motion. In condition 2, one of the motors detaches, leaving the endosome tethered to the microtubule but free to swivel. In condition 3, some regulatory factor [for example, Ndel1 (9)] halts processive motion of the motors, but they remain bound to the microtubule, restricting both translational and rotational motions. Conditions 4 and 5 illustrate two instances of direction reversal. In condition 4, the previously unbound kinesin binds and overpowers the dyneins in a tug-of-war. This manifests in an initial change in angle as the endosome rotates to relieve the torque of the kinesin binding, but the orientation is stable thereafter. In condition 5, the kinesin binds, whereas the dyneins unbind (8), and the endosome continues to rotate during subsequent motion. In condition 6, the previously unbound dynein becomes the leading motor exerting a torque on the endosome, causing an initial rotation as in condition 4. The new organization of the motor team causes a change in transport velocity.

Fig. 2. Validation of multipolarization dark-field imaging for orientation determination.

(A) Gold nanorods are excited identically in both two- and three-channel imaging, but their images are split in either two-component (ii) or three-component (iii) polarizations. Nanorods appear as diffraction-limited point spread functions. Ch, channel; PBS, polarizing beamsplitter; NPBS, nonpolarizing beamsplitter. (B) Calibration of azimuthal angle resolution for two-channel (i) and three-channel (ii) dark-field microscopy. Rotating the entire stage results in sinusoidal changes in intensity of individual nanorods (triangle data and line fit; i and ii, top). The calculated angle (i and ii, bottom, blue) matches well with the measured angle of the stage (gray). Error bars represent SD of the angle calculated for individual nanorods over a 75-frame movie. AU, arbitrary units.

Fig. 3. Axonal transport of gold nanorods in microfluidic culture.

(A) Neurons are plated close to the openings of microchannels into which they grow axons, which, therefore, have uniform directionality enabling unambiguous discrimination between dynein- and kinesin-mediated motion (scale bar, 100 μm). (B) WGA-nanorods are selectively added to the cell body or axon compartments where they are endocytosed and transported into the channels. Dark-field imaging is performed in the channels free from unbound and nonspecifically adsorbed nanorods. PDMS, polydimethylsiloxane. (C) Overlaying kymographs from 0 and π/2 polarization channels (green and magenta) show typical retrograde (i and movie S2) and anterograde (ii and movie S3) translational motion but also distinct rotational dynamics. Vertical scale bar, 5 μm; horizontal scale bar, 1 s. (D) Single-particle tracking of the endosome from (i) allows for precise angle (green) and position determination (blue).

Fig. 4. Gold nanorods as effective in vivo rotational probes.

(A) Nanorods attached to the plasma membrane (i) and supported bilayers (ii) show high rotational lability, whereas nanorods adsorbed to glass (iii) and WGA-nanorod-endosomes purified from neurons (iv) show much less rotational lability. (B) WGA-nanorod-endosomes are distinguished from free nanorods by colocalization with membrane staining FM1-43. (C) Cumulative distribution of σ from many nanorods (n = 268 for lipid bilayer, n = 69 for cell surface, n = 27 for glass surface, and n = 13 for endosome) confirms that there is minimal rotation of the nanorod with respect to the endosome. CDF, cumulative distribution function. (D) Example of an angle trace (top) converted into σ (bottom) with a long period of high rotational lability and a short one (arrows). Dashed gray line indicates σ = 0.044 above which is considered active rotation. (E) The duration of periods of increased rotational lability in processive retrograde trajectories shows that most are quite brief, and endosomes spend most of their time rotationally constrained (inset).

Fig. 5. Translational acceleration correlates with rotational dynamics.

(A) Velocity changes in retrograde-directed endosomes often coincided with angle changes; discrete velocity segments colored alternating black and blue with corresponding orientations colored dark and light green (movie S4). (B) Mean cross-correlation between absolute (top) or directional (bottom) translational acceleration and angular velocity in retrograde endosomes shows pronounced peak (blue) relative to time-scrambled control (red) around zero lag (error bars ± SEM by bootstrapping). (C and D) Similar correlation between translational acceleration and rotational velocity is evident in anterograde-directed endosomes, although with slightly lower correlation.

Fig. 6. Endosomes do not increase rotation inside pauses.

(A) Retrograde-directed (i) and anterograde-directed (ii) endosomes sometimes show translational pauses (directed motion is shown in blue; pauses are shown in black; corresponding orientations are shown in dark and light green, respectively). (B) Average rotational lability (σ) does not increase with pause duration for retrograde-directed (i) or anterograde-directed (ii) endosomes. It is also similar to mean σ during directed motion. Red bar is the median value for given pause length, and boxes represent 25th and 75th percentiles with black bars extending to minima and maxima. (C) Overall, rotational lability in pauses (black) closely matched directed motion (blue) but was more constrained than during reversals (red) for both net retrograde (top) and net anterograde (bottom) trajectories (±SEM by bootstrapping).

Fig. 7. Endosomes increase rotation following reversals.

(A) A representative endosome undergoing reversals clearly shows elevated rotation during reversals. Retrograde translocation is colored blue, reversals are colored red, and the corresponding angle points are colored dark greed and light green, respectively. (B) Distribution of change in mean σ from retrograde motion to pauses (blue) versus retrograde motion and reversals (red) in net retrograde trajectories. Histogram of the same distribution inset (error bars, 95% confidence interval by bootstrapping). (C) Alignment of reversals at zero time shows average σ (blue) increases significantly after reversal more than after nonreversal velocity changes (red) and random points (green) in the same set of trajectories (error bounds ± SEM).

Fig. 8. WGA-nanorod-endosomes align in direction of transport.

(A) The cumulative distribution of all processive retrograde (1472 trajectories; blue) and anterograde (231 trajectories; red) angles measured in three-channel imaging shows clear preference for alignment (zero angle) with microtubules. Error bars ± SEM from bootstrapping. (B) Neither absolute angle of the nanorod (top) nor rotational lability (bottom) is a strong function of velocity in retrograde-directed endosomes. (C) Rarely, endosomes show clear detachments from the microtubules (marked with red boxes) accompanied by much less constrained diffusion (movie S8). The alignment is much more uniformly distributed during the first pause as compared to the processive movement indicated by the dashed cyan box (distribution inset). This is evident in an x-y time projection plot (ii) where each frame in the trajectory is colored by its angle relative to the microtubule. (D) Retrograde-directed nanorods fully loaded with WGA show a high degree of alignment in two-channel microscopy (blue; n = 1736 endosomes). Reducing the number of WGA by blocking streptavidins with 10 times more biotin–bovine serum albumin (BSA) than nanorods leads to a reduced degree of alignment (black; n = 211). Further reducing the number of WGA (blocking with 1:60 nanorod/biotin-BSA) leads to even less alignment (green; n = 68). Shaded error bars represent ± SEM from bootstrapping. (E) Cumulative distribution of purified nanorod-containing endosomes (solid black line) also shows alignment with microtubules in vitro (n = 47 endosomes) compared with a uniform distribution (dashed gray line), although not as much as the median nanorod alignment during transport (blue line; same data as retrograde data from fig. S8). (F) Proposed mechanism of alignment featuring a direct linkage between ligand and microtubule through the endosomal membrane.