In conjunction with Foresight Update 53
Will Intracellular Medical Nanorobots Disrupt the Cytoskeleton (Part II)?
By Robert A. Freitas Jr.
Research Scientist, Zyvex Corp.
 |
| Robert A. Freitas Jr. |
Mechanical biocompatibility, or mechanocompatibility [1], is a critical issue in medical nanorobotics [1, 2]. For instance, active medical nanorobots maneuvering inside living cells could disturb or disrupt any of the many functions of the cytoskeleton. The cytoskeleton is the internal structural framework of a cell consisting of at least three types of filaments (microfilaments, microtubules, and intermediate filaments), forming a dynamic framework for maintaining cell shape and motion and allowing rapid changes in the three-dimensional structure of the cell ([2], Section 8.5.3.11).
Nanorobots could mechanically disrupt any of these functions during intracellular locomotion and manipulation of cell structures, if cytoskeletal/membrane links are disturbed. As already noted in Part I of this article, two classes of danger seem most significant. First, there is the risk of direct mechanical cytoskeletal reorganization, the subject of Part I (last time). Second, there is the risk of possible disruption of molecular motor-driven vesicular and organelle transport and related molecular motor diseases, the subject of Part II of this article. Three important examples of intracellular motor molecules are myosin (interdigitating myosin and actin filaments producing muscle contractions; [2], Section 6.3.4.2), dynein (drives motion of cilia; [2], Section 9.3.1.1), and kinesin (transport molecules carrying vesicles along microtubule tracks; [2], Section 9.4.6).
There are a variety of human disorders associated with dysfunction of cytoskeleton-based molecular motors, including, for instance, the motor-based diseases involving defective cellular myosin motors (e.g., implicated in hypertrophic cardiomyopathy [3]), spindle assembly- and function-related diseases [4], and other avenues for cellular malfunction. Cellular motors also participate in the self-organization of microtubule network structures [5]. But perhaps the most important function of molecular motors is intracellular vesicular transport, and most particularly axonal transport in neurons. Typically, organelles, vesicles and granules ~100 nm in diameter or larger are carried at a peak speed of up to ~2 microns/sec on the back of a 60-nm kinesin transport molecule ([2], Figure 9.32) that takes 8-nm ATP-powered steps along microtubule tracks running throughout the cell. (Mean unloaded kinesin motor speed is usually only 0.5-0.8 microns/sec.)
Conventional kinesin is a dimer of identical ~120 kD protein chains [6] with a diffusion coefficient D ~ 2 x 10-11 m2/sec [5]. The vesicle-attached kinesin motor molecule steps toward the plus-end of microtubules by converting the energy of ATP hydrolysis to mechanical work.
Could a vesicle be dislodged from its microtubule track after being bumped by a passing intracellular nanorobot? A kinesin molecular mechanical detachment force of Fdetach ~ 13 pN [8] requires that a 1-micron3 diamondoid nanorobot of mass mnano ~ 2 x 10-15 kg must impact and carry the cargo vesicle a distance Svesicle ~ 1 nm to detach it at a constant velocity of vnano >~ (2 Svesicle Fdetach / mnano)1/2 ~ 3500 micron/sec, well above the self-imposed 10 micron/sec intracellular locomotion speed limit ([1], Section 15.5.7.3.1) and even slightly exceeding the nanorobot instantaneous thermal velocity of ~2500 microns/sec in water at 37 °C ([2], Eqn. 3.3). The torsional stiffness of kinesin is so low that the molecule readily twists through more than 360° from its resting orientation [10], thus allowing the cargo to easily swivel out of the way of foreign impacting objects. And kinesin motors normally detach from a microtubule after a few seconds of travel anyway [6, 11]. Still, nanorobots should be able to exert mechanical forces well in excess of 13 pN, so care should be taken to minimize those motions and trajectories which are likely to produce kinesin detachment. Note that it has been found experimentally that intracellularly-placed microspheres (crude nanorobot analogues) experience enhanced diffusion over short time scales near the nucleus, possibly due to interactions with microtubule-associated motor proteins [12].
Could a nanorobot that has clamped or securely bound itself to one location on a microtubule prevent the passage of vesicular cargoes, causing vesicles to bunch up behind or to detach? And could kinesin motor molecules that encounter the obstruction be permanently damaged? Most likely a detached vesicle will reattach to a clear neighboring microtubule and continue its trek [13], or will reattach to the original microtubule downstream of the nanorobotic obstruction. In some cases organelles can attach to and move along multiple filaments simultaneously [14] using multiple motor molecules, potentially reducing the interference with forward motion from a single-filament blockage.
Coppin et al. [8] have carefully studied the behavior of kinesin molecules whose forward progress is mechanically constrained. Kinesin has a stall load of ~5 pN [7-9]. There is an increasing rate of dissociation with increasing load. Specifically, the dissociation rate is ~0.2/sec at 1 pN load, ~0.5/sec at 2 pN, ~1/sec at 3 pN, and ~2/sec at 5 pN load [8], rates which can be altered by the presence of microtubule-associated proteins [15]. However, super stall loads of 5-13 pN do not cause kinesin to walk backwards, “probably because of an irreversible transition in the mechanical cycle.” Rather, when super-stalled the kinesin motor most commonly takes a single backward movement and then dissociates (detaches) from the microtubule,* occasionally rebinding to the same microtubule (always at/below the stall load) and resuming its normal movement. This clearly demonstrates that the kinesin motor is still functional after being subjected to a dissociative induced stall. That is, a superload-induced detachment doesn’t “break” the biological motor. The conclusion is that similar mechanical interference by a nanorobot also should not damage a processive protein motor. Interestingly, a mutant form of kinesin with its ATP and microtubule binding sites decoupled has been found that binds so tightly to the microtubule that the motor cannot let go [16], crudely analogous to the case of a nanorobot that firmly grasps a microtubule for a period long in comparison to the timescale of kinesin procession.
* Alternating back and forth movements also are observed at super-stall [8]. The dissociation rate increases with load as long as the motor is moving (up to 2 Hz at 5 pN), but then becomes independent of load once the motor stalls — e.g., the stall time is 0.57 sec, representing a dissociation rate of ~1.8 Hz, for either spontaneous (~5 pN) or induced (~12 pN) stalls. Interestingly, forward loads induce the kinesin motor molecule to step faster under a wide range of ATP concentrations [8]. Forward loads of 5 pN increase velocity by +200% if ATP concentration is rate-limiting (5-40 µM) or by +50% if ATP is saturated (1 mM), but forward loads >5 pN cause forward velocity to drop off sharply [8]. A small carboxyl domain acts as a switch that turns the motor off when the kinesin motor is not bound to cargo [17].
A nanorobot ambulating along microtubules should endeavor to avoid applying lateral forces exceeding the kinesin detachment load of ~13 pN [8] which could have the effect of detaching associated vesicles as the nanorobot progresses — something like a tree limb being stripped of its leaves as it is pulled through a tight-fitting metal ring. Typically, the processive kinesin molecule only takes a few hundred steps before letting go [6, 11], so an occasional early detachment, by itself, should not induce cellular dysfunction. As long as the processive motor protein is not physically damaged, most of the detached vesicles should reattach and continue their journey after the nanorobot has passed by. The minimum spacing (maximum density) of kinesin motors along a microtubule is an axial repeat of 8 nm and experiments with isolated microtubules gliding on kinesin-coated glass find optimal motility at a 47 nm separation between kinesins [18]. But the vesicular transport of 100-2000 nm diameter organelles in cyto implies that propulsive kinesin contacts are normally spaced at least 0.1-2 microns apart along the microtubule tracks. This leaves plenty of unoccupied foothold space to allow nanorobots to avoid disrupting processive motor proteins already in transit.
Coordinated groups of in cyto medical nanorobots should avoid inadvertently corralling or bulldozing large numbers of vesicles or motor molecules into relatively small volumes within the cell, as such increases in motor molecule concentration could increase the local microtubule polymerization rate [5]. Analogously, in motor neuron diseases where vesicular transport is blocked by massive localized accumulations of kinesin molecules, the blockage can produce large axonal swellings in the motor neurons in human spinal cords and can disturb the machinery for anterograde fast axonal transport [19].
Intracellular traffic jams involving repositioned vesicles and organelles [20] appear to be initiated by the accumulation of stalled kinesin cargoes and are most commonly reported in neurons [21-23] where their effects are most serious. For example, axonal organelles transported by kinesin molecules that stall can cause organelle jams that disrupt retrograde as well as anterograde fast axonal transport, leading to defective action potentials, dystrophic terminals, reduced transmitter secretion and progressive distal paralysis that parallels the pathologies of motor diseases such as amyotrophic lateral sclerosis [23]. Stretch injury to axonal cytoskeleton resulting in major loss of microtubules disrupts fast axonal transport resulting in focal accumulation of membranous organelles and axonal swellings [22], and a chemically-created microtubule-free region can serve as a trap that causes axonally transported particles to accumulate into a swollen region [21]. Nanorobots should avoid creating such regions within the cell.
Similar considerations also apply during nanorobotic intranuclear operations, given the presence of myosin-based motors, RNA polymerase motors, and other motor molecules inside the cell nucleus.
Copyright 2003 Robert A. Freitas Jr. All Rights Reserved
References
1. R.A. Freitas Jr., Nanomedicine, Volume IIA: Biocompatibility, Landes Bioscience, Georgetown, TX, 2003; http://www.nanomedicine.com
2. R.A. Freitas Jr., Nanomedicine, Volume I: Basic Capabilities, Landes Bioscience, Georgetown, TX, 1999; http://www.nanomedicine.com
3. I. Rayment et al., “Structural interpretation of the mutations in the beta-cardiac myosin that have been implanted in familial hypertrophic cardiomyopathy,” Proc. Natl. Acad. Sci. (USA) 92(25 April 1995):3864-3868.
4. V. Mountain, D.A. Compton, “Dissecting the role of molecular motors in the mitotic spindle,” Anat. Rec. 261(15 February 2000):14-24.
5. T. Surrey et al., “Physical properties determining self-organization of motors and microtubules,” Science 292(11 May 2001):1167-1171.
6. S. Rice et al., “A structural change in the kinesin motor protein that drives motility,” Nature 402(16 December 1999):778-784.
7. K. Svoboda et al., “Direct observation of kinesin stepping by optical trapping interferometry,” Nature 365(21 October 1993):721-727.
8. C.M. Coppin et al., “The load dependence of kinesin’s mechanical cycle,” Proc. Natl. Acad. Sci. (USA) 94(5 August 1997):8539-8544.
9. M.E. Fisher, A.B. Kolomeisky, “The force exerted by a molecular motor,” Proc. Natl. Acad. Sci. (USA) 96(8 June 1999):6597-6602.
10. A.J. Hunt, J. Howard, “Kinesin swivels to permit microtubule movement in any direction,” Proc. Natl. Acad. Sci. (USA) 90(15 December 1993):11653-11657.
11. W.O. Hancock, J. Howard, “Kinesin’s processivity results from mechanical and chemical coordination between the ATP hydrolysis cycles of the two motor domains,” Proc. Natl. Acad. Sci. (USA) 96(9 November 1999):13147-13152.
12. A. Caspi et al., “Enhanced diffusion in active intracellular transport,” Phys. Rev. Lett. 85(25 December 2000):5655-5658.
13. Y. Wada et al., “Evidence for a novel affinity mechanism of motor-assisted transport along microtubules,” Mol. Biol. Cell 11(January 2000):161-169.
14. R.D. Vale et al., “Movement of organelles along filaments dissociated from the axoplasm of the squid giant axon,” Cell 40(February 1985):449-454.
15. B. Trinczek et al., “Tau regulates the attachment/detachment but not the speed of motors in microtubule-dependent transport of single vesicles and organelles,” J. Cell Sci. 112(July 1999):2355-2367.
16. H. Song, S.A. Endow, “Decoupling of nucleotide- and microtubule-binding sites in a kinesin mutant,” Nature 396(10 December 1998):587-590.
17. D.L. Coy et al., “Kinesin’s tail domain is an inhibitory regulator of the motor domain,” Nat. Cell Biol. 1(September 1999):288-292.
18. K.J. Bohm et al., “Speeding up kinesin-driven microtubule gliding in vitro by variation of cofactor composition and physicochemical parameters,” Cell. Biol. Int. 24(2000):335-341.
19. I. Toyoshima et al., “Kinesin and cytoplasmic dynein in spinal spheroids with motor neuron disease,” J. Neurol. Sci. 159(15 July 1998):38-44.
20. R.A. Spritz, “Multi-organelle disorders of pigmentation: intracellular traffic jams in mammals, flies and yeast,” Trends Genet. 15(September 1999):337-340.
21. H. Horie et al., “Effects of disruption of microtubules on translocation of particles and morphology in tissue cultured neurites,” Brain Res. 288(12 December 1983):85-93.
22. W.L. Maxwell, “Histopathological changes at central nodes of Ranvier after stretch-injury,” Microsc. Res. Tech. 34(15 August 1996):522-535.
23. D.D. Hurd, W.M. Saxton, “Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila,” Genetics 144(November 1996):1075-1085.
More on the medical applications of nanotechnology at: http://www.foresight.org/Nanomedicine/index.html
Also, please check out http://www.nanomedicine.com/. Robert Freitas has acquired ownership of the domain, and has put up a cleaner and very extensively internally-linked version of Nanomedicine Vol. I at http://www.nanomedicine.com/NMI.htm
IMM would appreciate learning your thoughts on the above article.