Superior cervical ganglion

Superior cervical ganglion DEFAULT

Superior cervical ganglion

The superior cervical ganglion (SCG) is part of the autonomic nervous system (ANS), more specifically it is part of the sympathetic nervous system, a division of the ANS most commonly associated with the fight or flight response. The ANS is composed of pathways that lead to and from ganglia, groups of nerve cells. A ganglion allows a large amount of divergence in a neuronal pathway and also enables a more localized circuitry for control of the innervated targets.[1] The SCG is the only ganglion in the sympathetic nervous system that innervates the head and neck. It is the largest and most rostral (superior) of the three cervical ganglia. The SCG innervates many organs, glands and parts of the carotid system in the head.

Structure[edit]

Location[edit]

The SCG is located opposite the second and third cervical vertebrae. It lies deep to the sheath of the internal carotid artery and internal jugular vein, and anterior to the Longus capitis muscle. The SCG contains neurons that supply sympathetic innervation to a number of target organs within the head.

The SCG also contributes to the cervical plexus. The cervical plexus is formed from a unification of the anterior divisions of the upper four cervical nerves. Each receives a gray ramus communicans from the superior cervical ganglion of the sympathetic trunk.[2]

Morphology and physiology and its neurons[edit]

The superior cervical ganglion is a reddish-gray color, and usually shaped like a spindle with tapering ends. Sometimes the SCG is broad and flattened, and occasionally constricted at intervals. It formed by the coalescence of four ganglia, corresponding to the upper four cervical nerves, C1-C4. The bodies of these preganglionic sympathetic neurons are specifically located in the lateral horn of the spinal cord. These preganglionic neurons then enter the SCG and synapse with the postganglionic neurons that leave the rostral end of the SCG and innervate target organs of the head.

There are a number of neuron types in the SCG ranging from low threshold to high threshold neurons. The neurons with a low threshold have faster action potential firing rate, while the high threshold neurons have a slow firing rate.[3] Another distinction between SCG neuron types is made via immunostaining. Immunostaining allows the classification of SCG neurons as either positive or negative for neuropeptide Y (NPY), which is found in a subgroup of high-threshold neurons.[3] Low threshold, NPY-negative neurons are secretomotor neurons, innervating salivary glands. High threshold, NPY-negative neurons are vasomotor neurons, innervating blood vessels. High threshold, NPY-positive neurons are vasoconstrictor neurons, which innervate the iris and pineal gland.

Innervation[edit]

The SCG receives input from the ciliospinal center. The ciliospinal center is located between the C8 and T1 regions of the spinal cord within the intermediolateral column. The preganglionic fibers that innervate the SCG are the thoracic spinal nerves, which extend from the T1-T8 region of the ciliospinal center. These nerves enter the SCG through the cervical sympathetic nerve. A mature preganglionic axon can innervate anywhere from SCG cells.[4] Postganglionic fibers then leave the SCG via the internal carotid nerve and the external carotid nerve. This pathway of SCG innervation is shown through stimulation of the cervical sympathetic nerve, which invokes action potentials in both the external and internal carotid nerves.[5] These postganglionic fibers shift from multiple axon innervation of their targets to less profound multiple axon innervation or single axon innervation as the SCG neurons mature during postnatal development.[6]

Function[edit]

Sympathetic nervous system[edit]

The SCG provides sympathetic innervation to structures within the head, including the pineal gland, the blood vessels in the cranial muscles and the brain, the choroid plexus, the eyes, the lacrimal glands, the carotid body, the salivary glands, and the thyroid gland.[1]

Pineal gland[edit]

The postganglionic axons of the SCG innervate the pineal gland and are involved in Circadian rhythm.[7] This connection regulates production of the hormone melatonin, which regulates sleep and wake cycles, however the influence of SCG neuron innervation of the pineal gland is not fully understood.[8]

Carotid body[edit]

The postganglionic axons of the SCG innervate the internal carotid artery and form the internal carotid plexus. The internal carotid plexus carries the postganglionic axons of the SCG to the eye, lacrimal gland, mucous membranes of the mouth, nose, and pharynx, and numerous blood-vessels in the head.

The eye[edit]

The postganglionic axons of the Superior cervical ganglion innervate the eye and lacrimal gland and cause vasoconstriction of the iris and sclera, pupillary dilation, widening of the palpebral fissure, and the reduced production of tears.[9] These responses are important during Fight-or-flight response of the ANS. Dilation of the pupils allows for an increased clarity in vision, and inhibition of the lacrimal gland stops tear production allowing for unimpaired vision and redirection of energy elsewhere.

Blood vessels of the skin[edit]

The postganglionic axons of the SCG innervate blood vessels in the skin and cause the vessels to constrict. Constriction of the blood vessels causes a decrease in blood flow to the skin leading to paling of the skin and retention of body heat. This plays into the fight-or-flight response, decreasing blood flow to facial skin and redirecting the blood to more important areas like the blood vessels of muscles.

Vestibular system[edit]

The SCG is connected with vestibular structures, including the neuroepithelium of the semicircular canals and otolith organs, providing a conceivable substrate for modulation of vestibulo-sympathetic reflexes.

Clinical significance[edit]

Horner's syndrome[edit]

Horner's syndrome is a disorder resulting from damage to the sympathetic autonomic nervous pathway in the head. Damage to the SCG, part of this system, often results in Horner's syndrome. Damage to the T1-T3 regions of the spinal cord is responsible for drooping of the eyelids (ptosis), constriction of the pupil (miosis), and sinking of the eyeball (apparent Enophthalmos; not truly sunken, just appears so because of the drooping eyelid).[7] Lesion or significant damage to the SCG results in a third order neuron disorder (see Horner's Syndrome: Pathophysiology).

Familial dysautonomia[edit]

Familial dysautonomia is a genetic disorder characterized by abnormalities of sensory and sympathetic neurons. The SCG is significantly affected by this loss of neurons and may be responsible for some of the resulting symptoms. In post-mortem studies the SCG is, on average, one-third of normal size and has only 12 percent of the normal number of neurons.[10] Defects in the genetic coding for NGF, which result in less functional, abnormally structured NGF, may be the molecular cause of familial dysautonomia.[11] NGF is necessary for survival of some neurons so loss of NGF function could be the cause for neuronal death in the SCG.

History[edit]

Reinnervation[edit]

In the late 19th century, John Langley discovered that the superior cervical ganglion is topographically organized. When certain areas of the superior cervical ganglion were stimulated, a reflex occurred in specified regions of the head. His findings showed that preganglionic neurons innervate specific postganglionic neurons.[6][12] In his further studies of the superior cervical ganglion, Langley discovered that the superior cervical ganglion is regenerative. Langley severed the SCG above the T1 portion, causing a loss of reflexes. When left to their own accord, the fibers reinnervated the SCG and the initial autonomic reflexes were recovered, though there was limited recovery of pineal gland function.[13] When Langley severed the connections between the SCG and the T1–T5 region of the spinal cord and replaced the SCG with a different one, the SCG was still innervated the same portion of the spinal cord as before. When he replaced the SCG with a T5 ganglion, the ganglion tended to be innervated by the posterior portion of the spinal cord (T4–T8). The replacement of the original SCG with either a different one or a T5 ganglion supported Langley's theory of topographic specificity of the SCG.

Research[edit]

Ganglia of the peripheral autonomic nervous system are commonly used to study synaptic connections. These ganglia are studied as synaptic connections show many similarities to the central nervous system (CNS) and are also relatively accessible. They are easier to study than the CNS since they have the ability to regrow, which neurons in the CNS do not have. The SCG is frequently used in these studies being one of the larger ganglia.[14] Today, neuroscientists are studying topics on the SCG such as survival and neurite outgrowth of SCG neurons, neuroendocrine aspects of the SCG, and structure and pathways of the SCG. These studies are usually performed on rats, guinea-pigs, and rabbits.

Historical contributions[edit]

  • E. Rubin studied the development of the SCG in fetal rats.[15] Research on the development of nerves in the SCG has implications for the general development of the nervous system.
  • The effects of age on dendritic arborisation of sympathetic neurons has been studied in the SCG of rats. Findings have shown that there is significant dendritic growth in the SCG of young rats but none in aged rats. In aged rats, it was found, that there was a reduction in the number of dendrites.[16]
  • SCG cells were used to study nerve growth factor (NGF) and its ability to direct growth of neurons. Results showed that NGF did have this directing, or tropic, effect on neurons, guiding the direction of their growth.[17]

Additional images[edit]

  • The right sympathetic chain and its connections with the thoracic, abdominal, and pelvic plexuses.

  • Superior cervical ganglion

  • Sympathetic connections of the ciliary and superior cervical ganglia.

  • The position and relation of the esophagus in the cervical region and in the posterior mediastinum. Seen from behind.

  • The Sympathetic Trunk and SCG innervation of target organs in the head.

References[edit]

Public domainThis article incorporates text in the public domain from page of&#;the 20th edition ofGray's Anatomy()

  1. ^ abMichael J. Zigmond, ed. (). Fundamental neuroscience (2&#;ed.). San Diego: Acad. Press. pp.&#;– ISBN&#;.
  2. ^Henry Gray. Anatomy of the Human Body. 20th ed. Philadelphia: Lea & Febiger, New York: Bartleby.com, http://www.bartleby.com//html. Accessed July 9,
  3. ^ abLi, Chen; Horn, John P. (). "Physiological classification of sympathetic neurons in the rat superior cervical ganglion". Journal of Neurophysiology. 95 (1): – doi/jn PMID&#;
  4. ^Purves, D; Wigston, DJ (January ). "Neural units in the superior cervical ganglion of the guinea-pig". The Journal of Physiology. (1): – doi/jphysiolsp PMC&#; PMID&#;
  5. ^Purnyn, H..; Rikhalsky, O.; Fedulova, S.; Veslovsky, N. (). "Transmission Pathways in the Rat Superior Cervical Ganglion". Neurophysiology. 39 (4–5): – doi/s S2CID&#;
  6. ^ abPurves, Dale; Lichtman, Jeff W. (). Development of the Nervous System. Sunderland, Mass.: Sinauer Associates. pp.&#;– ISBN&#;.
  7. ^ abPurves, Dale (). Neuroscience (5&#;ed.). Sunderland, Mass.: Sinauer. p.&#; ISBN&#;.
  8. ^Photoperiodism, melatonin, and the pineal. London: Pitman Publishing Ltd. p.&#;
  9. ^Lichtman, Jeff W.; Purves, Dale; Yip, Joseph W. (). "On the purpose of selective innervation of guinea-pig superior cervical ganglion cells". Journal of Physiology. (1): 69– doi/jphysiolsp PMC&#; PMID&#;
  10. ^Pearson, J; Brandeis, L; Goldstein, M (5 October ). "Tyrosine hydroxylase immunoreactivity in familial dysautonomia". Science. (): 71– BibcodeSciP. doi/science PMID&#;
  11. ^Schwartz, JP; Breakefield, XO (February ). "Altered nerve growth factor in fibroblasts from patients with familial dysautonomia". Proceedings of the National Academy of Sciences of the United States of America. 77 (2): –8. BibcodePNASS. doi/pnas PMC&#; PMID&#;
  12. ^Sanes, Dan H.; Reh, Thomas A.; Harris, William A. (). Principles of neural development. San Diego, CA: Academic Press. pp.&#;– ISBN&#;.
  13. ^Lingappa, Jaisri R.; Zigmond, Richard E. (). "Limited Recovery of Pineal Function after Regeneration of Preganglionic Sympathetic Axons:Evidence for Loss of Ganglionic Synaptic Specificity". The Journal of Neuroscience. 33 (11): – doi/JNEUROSCI PMC&#; PMID&#;
  14. ^Purves, D; Lichtman, JW (October ). "Formation and maintenance of synaptic connections in autonomic ganglia". Physiological Reviews. 58 (4): – doi/physrev PMID&#;
  15. ^Rubin, E (March ). "Development of the rat superior cervical ganglion: ganglion cell maturation". The Journal of Neuroscience. 5 (3): – doi/jneurosci PMC&#; PMID&#;
  16. ^Andrews, TJ; Li, D; Halliwell, J; Cowen, T (February ). "The effect of age on dendrites in the rat superior cervical ganglion". Journal of Anatomy. (1): –7. PMC&#; PMID&#;
  17. ^Campenot, RB (). "Local control of neurite development by nerve growth factor". Proc Natl Acad Sci U S A. 74 (10): –9. BibcodePNASC. doi/pnas PMC&#; PMID&#;

External links[edit]

Sours: https://en.wikipedia.org/wiki/Superior_cervical_ganglion

Superior cervical ganglion

Template:Infobox NerveEditor-In-Chief:C. Michael Gibson, M.S., M.D.[1]


The superior cervical ganglion (SCG), the largest of the cervical ganglia, is placed opposite the second and third cervical vertebræ. It contains neurons that supply sympathetic innervation to the face.

It is of a reddish-gray color, and usually fusiform in shape; sometimes broad and flattened, and occasionally constricted at intervals; it is believed to be formed by the coalescence of four ganglia, corresponding to the upper four cervical nerves.

It is in relation, in front, with the sheath of the internal carotid artery and internal jugular vein; behind, with the Longus capitis muscle.

It receives input from the ciliospinal center.

Additional images

  • The right sympathetic chain and its connections with the thoracic, abdominal, and pelvic plexuses.

  • Diagram of efferent sympathetic nervous system.

  • Sympathetic connections of the ciliary and superior cervical ganglia.

  • The position and relation of the esophagus in the cervical region and in the posterior mediastinum. Seen from behind.

External links


Template:Gray'sTemplate:Autonomic

Template:WikiDoc Sources

de:Ganglion cervicale superius

Sours: https://www.wikidoc.org/index.php/Superior_cervical_ganglion
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Neuroanatomy, Superior Cervical Ganglion

Introduction

The superior cervical ganglia are involved in the autonomic nervous system. They are involved explicitly with sympathetic efferent innervation, particularly to the face and head. The superior cervical ganglion is the most superior ganglion of the sympathetic chain, bilaterally. It is the largest of the three ganglia of the cervical sympathetic trunk, the other two being the stellate and middle cervical ganglion.[1] The superior cervical ganglia are approximately at the levels of the second cervical vertebra (C2) and the third cervical vertebra (C3), bilaterally. They typically lie lateral to the longus colli muscles, bilaterally. The superior cervical ganglia usually lie anteromedial or medial to their respective internal carotid arteries.[1][2] They are considered a landmark for the sympathetic trunk and typically provide sympathetic innervation to the head and neck.[3][4]

Structure and Function

The superior cervical ganglia are made up of the paravertebral ganglia of cervical vertebral levels one through four (C1-C4).[3][4] The superior cervical ganglia are oval shaped, cylinder-like, and elongated. They contain neurons.[1][5][6] They receive presynaptic signals from the sympathetic trunk. Postsynaptic signals then convey information from the superior cervical ganglia to targets in the head and neck.[2]

Embryology

The superior cervical ganglia derive from neural crest cells. Neural crest cells come from the ectoderm in embryological development. Many structures of the head, neck, face, and cardiovascular system also originate from the neural crest cells.[5][7]

Blood Supply and Lymphatics

Branches from the ascending pharyngeal artery, coming off of the external carotid artery, provide blood supply to the superior cervical ganglia. Venous drainage is through the internal jugular vein through direct posterior branches. There is a noted deficiency in gross blood supply at the junction of the upper one third and lower two-thirds of the neck; this is important for surgical considerations mentioned in the later part of this activity.[8]

The superior cervical ganglia are anterior to the retropharyngeal lymph nodes.[1][2]

Nerves

As previously described, the superior cervical ganglia are made up of the paravertebral ganglia of cervical vertebral levels one through four (C1-C4).[3][4] The superior cervical ganglia themselves contain neurons that supply sympathetic innervation to the head and neck.[5][6] The superior cervical ganglia provide postsynaptic fibers via the deep petrosal nerve to provide vasoconstriction to the lacrimal gland.[9] The superior cervical ganglia also supply gray rami communicantes to the ventral rami in the cervical plexus.[10] The superior cervical ganglia connect to the next closest ganglia, the middle cervical ganglia, inferiorly via the sympathetic trunk.[4]

Muscles

The superior cervical ganglia provide sympathetic innervation.[10] Their location is lateral to the longus colli muscles.[1][2]

Physiologic Variants

The superior cervical ganglia are typically present at cervical vertebral levels one and two (C1 and C2). However, they have been found between the cervical vertebral levels one through five. (C1-C5).[1] The ranges of the length of the superior cervical ganglia are from 10 to 30mm. The width of the superior cervical ganglia ranges from 5 to 8mm.[1]

Branches coming off of the superior cervical ganglia include the:

  • Pharyngeal branch

  • The communicating branch of the cervical nerve

  • The internal carotid branch

  • The communicating branch of the pharyngeal mucosa

  • The communicating branch of the vagus nerve

  • The communicating branch of the superior laryngeal nerve

  • The laryngeal branch

  • The communicating branch of the internal jugular vein

There are reports of up to seven different types of branch combinations coming off of the superior cervical ganglia.[2]

It is crucial to recognize that these are common variants of branches coming off of the superior cervical ganglia as damaging these may lead to consequences such as the development of Horner syndrome as described later in this activity.[1]

Surgical Considerations

It is important to avoid this the superior cervical ganglia during open anterior neck surgeries. When the superior cervical ganglia are damaged, Horner syndrome can develop. Horner syndrome comes from tension on the sympathetic trunk during anterior neck surgeries such as anterior cervical disc fusions. The tension causes damage to the fibers along the sympathetic trunk. More about Horner syndrome is described later in this activity.[1][4][11] Knowing the anatomy and landmarks regarding the neck helps to avoid damaging important neurovasculature in the area.[1]

Anterior surgical procedures to the neck area such as anterior cervical discectomies can cause a disruption of the blood supply to the superior cervical ganglia, causing ischemic damage to the ganglia. This ischemic damage can also cause symptoms similar to Horner syndrome.[7]

To prevent damage to the sympathetic ganglia during an anterior approach to the cervical spine, it is always advisable to create longus colli gutter and then place the retractors within the same, thereby minimizing the risk of inadvertent injury to the ganglia that lie lateral to the muscle.

Clinical Significance

The superior cervical ganglia can be a site for locally injected anesthetics such as opioids, which is useful for neuropathic pain management of the face.[1][2] The superior cervical ganglia are the main source of sympathetic innervation to the head and face. Buprenorphine has been specifically utilized in these ganglionic local opioid analgesia injections to provide pain relief to patients suffering from neuropathic facial pain stemming from atypical facial pain, postherpetic neuralgia, and trigeminal neuralgia.[12] Understanding the anatomic positioning of the superior cervical ganglia allows this injection to be performed safely by avoiding important structures such as the internal carotid arteries which lie close to the superior cervical ganglia.[1][2][12]. Ultrasound guidance can be utilized to visualize the superior cervical ganglia during this procedure properly.[12]

Damage to the superior cervical ganglia can cause a postganglionic pattern of Horner syndrome. The symptoms of Horner syndrome include miosis, facial anhidrosis, and partial ptosis; this is due to the superior cervical ganglia's role in providing sympathetic innervation to these targets.[4][11] These symptoms arise from the disruption of the sympathetic nervous system, which includes superior cervical ganglia. Care is necessary during superior cervical ganglia blocks as well as head and neck procedures to prevent Horner syndrome from occurring.[11]

Other Issues

The superior cervical ganglion can appear as pathologic retropharyngeal lymph nodes on imaging. The superior cervical ganglia are anterior to the retropharyngeal lymph nodes.[1][2] The proximity of the superior cervical ganglia to the retropharyngeal lymph nodes can make the staging of neck neoplasms difficult. Nasopharyngeal cancer, thyroid cancer, as well as oropharyngeal cancer, typically metastasizes to the retropharyngeal lymph nodes, therefore delineating them from the superior cervical ganglia is very important.[1][2][3] Magnetic resonance imaging can be utilized to delineate the difference between pathologic retropharyngeal lymph nodes from the superior ganglia.[1][2]

The Sympathetic Nerves, Sympathetic connections of the ciliary and superior cervical ganglia

Figure

The Sympathetic Nerves, Sympathetic connections of the ciliary and superior cervical ganglia. Contributed by Gray's Anatomy Plates

The Sympathetic Nerves, Sympathetic connections of the sphenopalatine and superior cervical ganglia

Figure

The Sympathetic Nerves, Sympathetic connections of the sphenopalatine and superior cervical ganglia. Contributed by Gray's Anatomy Plates

The Sympathetic Nerves, Sympathetic connections of the submaxillary and superior cervical ganglia

Figure

The Sympathetic Nerves, Sympathetic connections of the submaxillary and superior cervical ganglia. Contributed by Gray's Anatomy Plates

The Cervical Portion of the Sympathetic System, Diagram of the cervical sympathetic

Figure

The Cervical Portion of the Sympathetic System, Diagram of the cervical sympathetic. Contributed by Gray's Anatomy Plates

The Cervical Portion of the Sympathetic System, Plan of right sympathetic cord and splanchnic nerves

Figure

The Cervical Portion of the Sympathetic System, Plan of right sympathetic cord and splanchnic nerves. Contributed by Gray's Anatomy Plates

References

1.

Yokota H, Mukai H, Hattori S, Yamada K, Anzai Y, Uno T. MR Imaging of the Superior Cervical Ganglion and Inferior Ganglion of the Vagus Nerve: Structures That Can Mimic Pathologic Retropharyngeal Lymph Nodes. AJNR Am J Neuroradiol. Jan;39(1) [PMC free article: PMC] [PubMed: ]

2.

Mitsuoka K, Kikutani T, Sato I. Morphological relationship between the superior cervical ganglion and cervical nerves in Japanese cadaver donors. Brain Behav. Feb;7(2):e [PMC free article: PMC] [PubMed: ]

3.

Loke SC, Karandikar A, Ravanelli M, Farina D, Goh JP, Ling EA, Maroldi R, Tan TY. Superior cervical ganglion mimicking retropharyngeal adenopathy in head and neck cancer patients: MRI features with anatomic, histologic, and surgical correlation. Neuroradiology. Jan;58(1) [PubMed: ]

4.

Fazliogullari Z, Kilic C, Karabulut AK, Yazar F. A morphometric analysis of the superior cervical ganglion and its surrounding structures. Surg Radiol Anat. Apr;38(3) [PubMed: ]

5.

Moriyama H, Shimada K, Goto N. Morphometric analysis of neurons in ganglia: geniculate, submandibular, cervical spinal and superior cervical. Okajimas Folia Anat Jpn. Oct;72(4) [PubMed: ]

6.

Rusu MC, Pop F. The anatomy of the sympathetic pathway through the pterygopalatine fossa in humans. Ann Anat. Feb 20;(1) [PubMed: ]

7.

Kameda Y, Saitoh T, Nemoto N, Katoh T, Iseki S. Hes1 is required for the development of the superior cervical ganglion of sympathetic trunk and the carotid body. Dev Dyn. Aug;(8) [PubMed: ]

8.

Tubbs RS, Salter G, Wellons JC, Oakes WJ. Blood supply of the human cervical sympathetic chain and ganglia. Eur J Morphol. Dec;40(5) [PubMed: ]

9.

Goosmann MM, Dalvin M. StatPearls [Internet]. StatPearls Publishing; Treasure Island (FL): Aug 13, Anatomy, Head and Neck, Deep Petrosal Nerve. [PubMed: ]

Waxenbaum JA, Reddy V, Bordoni B. StatPearls [Internet]. StatPearls Publishing; Treasure Island (FL): Feb 7, Anatomy, Head and Neck, Cervical Nerves. [PubMed: ]

Khan Z, Bollu PC. StatPearls [Internet]. StatPearls Publishing; Treasure Island (FL): May 4, Horner Syndrome. [PubMed: ]

Siegenthaler A, Haug M, Eichenberger U, Suter MR, Moriggl B. Block of the superior cervical ganglion, description of a novel ultrasound-guided technique in human cadavers. Pain Med. May;14(5) [PubMed: ]

Sours: https://www.ncbi.nlm.nih.gov/books/NBK/
Cervical portion of sympathetic trunk

Loss of Cervical Sympathetic Chain Input to the Superior Cervical Ganglia Affects the Ventilatory Responses to Hypoxic Challenge in Freely-Moving C57BL6 Mice

Introduction

Pre-ganglionic sympathetic neurons emanating from the thoracic spinal cord (T1&#x;T4) course in the left and right cervical sympathetic chains (CSC) and terminate on the cell bodies of post-ganglionic sympathetic neurons in the ipsilateral superior cervical ganglia (SCG) (Rando et al., ; Tang et al., a,b,c; Llewellyn-Smith et al., ). The majority of post-ganglionic cells leave the SCG via the internal and external carotid nerves (Bowers and Zigmond, ; Buller and Bolter, ; Asamoto, ; Savastano et al., ), with the ganglioglomerular nerve (GGN) branching from the external carotid nerve to innervate glomus cells, chemoafferent nerve terminals, and vasculature within the carotid body (Biscoe and Purves, ; Zapata et al., ; Bowers and Zigmond, ; McDonald and Mitchell, ; McDonald, ; Verna et al., ; Torrealba and Claps, ; Ichikawa, ; Asamoto, ; Savastano et al., ), and baroafferent nerve terminals within the carotid sinus (Floyd and Neil, ; Rees, ; Bolter and Ledsome, ; Felder et al., ; Buller and Bolter, ).

The projections of the SCG to a series of inter-related structures, such as the carotid body and carotid sinus, upper airway and tongue (Flett and Bell, ; Kummer et al., ; O&#x;Halloran et al., , ; Hisa et al., ; Wang and Chiou, ; Oh et al., ), and nuclei within the hypothalamus and brainstem, including the nucleus tractus solitarius (nTS) (Cardinali et al., a,b, ; Gallardo et al., ; Saavedra, ; Wiberg and Widenfalk, ; Westerhaus and Loewy, ; Esquifino et al., ; Hughes-Davis et al., ; Mathew, ), provide evidence that the SCG is a vital integrative structure regulating cardiorespiratory function. Previous studies have shown that electrical stimulation of the CSC decreases arterial blood pressure and upper airway resistance in rats (O&#x;Halloran et al., , ). Moreover, there is substantial (and conflicting) evidence as to the ability of sympathetic innervation to the carotid body to influence resting activity of glomus cells and chemoafferents within the carotid sinus nerve (CSN), and modulate changes in activity during hypoxic gas challenge (HXC) (Prabhakar, ). For instance, CSC and GGN activity increases during hypoxic challenge, suggesting the release of neurotransmitters, such as norepinephrine, dopamine and neuropeptide Y (Lahiri et al., ; Matsumoto et al., , , Yokoyama et al., ), and conversely activation of the GGN decreases the hypoxic response of chemosensors within the cat carotid body (McQueen et al., ). A variety of responses have also been reported upon application of neurotransmitters released by CSC and GGN nerve terminals (e.g., norepinephrine, dopamine and neuropeptide Y) to in vivo and in vitro carotid body preparations, including (1) a biphasic pattern consisting of initial brief bursts in CSN activity and then long-lasting inhibition (Bisgard et al., ), (2) a biphasic pattern consisting of initial brief decreases in CSN activity and then long-lasting excitation (Matsumoto et al., ), (3) indirect excitation of carotid body glomus cells via constriction of arteriolar blood flow in the carotid body (Potter and McCloskey, ; Yokoyama et al., ), (4) direct activation of glomus cells and/or chemosensory afferents (Matsumoto et al., ; Lahiri et al., ; Milsom and Sadig, ; Heinert et al., ; Pang et al., ), and (5) direct inhibitory action of glomus cells and/or chemosensory afferents (Zapata et al., ; Zapata, ; Llados and Zapata, ; Mills et al., ; Folgering et al., ; Kou et al., ; Pizarro et al., ; Bisgard et al., ; Prabhakar et al., ; Ryan et al., ; Almaraz et al., ; Overholt and Prabhakar, ).

Wild-type and genetically-engineered mice are widely used to understand the mechanisms by which hypoxic challenges elicit carotid body-dependent and -independent ventilatory responses (He et al., , , ; Kline and Prabhakar, ; Pérez-García et al., ; Kline et al., ; Pichard et al., ; Gao et al., ; Wang et al., ; Ortega-Sáenz et al., ; Peng et al., ; Prabhakar et al., ). The morphology, neurophysiology and neuropharmacology of the mouse CSC-SCG has received considerable investigation over the years (Black et al., ; Yokota and Yamauchi, ; Banks and Walter, ; Inoue, ; Lewis and Burton, ; Forehand, ; Kidd and Heath, ; Gibbins, ; Kasa et al., ; Little and Heath, ; Jobling and Gibbins, ; El-Fadaly and Kummer, ; David et al., ; Cadaveira-Mosquera et al., ; Pashai et al., ; Alberola-Die et al., ; Liu and Bean, ; Martinez-Pinna et al., ; Mitsuoka et al., ; Feldman-Goriachnik and Hanani, ; Simeone et al., ; Rivas-Ramírez et al., ). Mouse tissues that receive post-ganglionic projections from the SCG have also been heavily investigated (Krieger et al., ; García et al., ; Kawaja and Crutcher, ; Maklad et al., ; Pankevich et al., a,b; Ivanusic et al., ; Karlsen et al., ; Lindborg et al., ; Ziegler et al., ; Teshima et al., ). Nonetheless, there is no direct evidence that the CSC-SCG complex innervates the carotid bodies of mice, and this is likely on the basis of the presence of sympathetic (i.e., tyrosine-hydroxylase-positive) nerve terminals and adrenergic receptors in these structures (Prieto-Lloret et al., ; Roux et al., ; Kåhlin et al., ; Chai et al., ).

Patients with chronic T1&#x;T4 spinal cord injury present with cardiorespiratory disturbances consistent with diminished activity of the CSC-SCG complex (DiMarco et al., ; Heutink et al., ; Sankari et al., , ; Berlowitz et al., ; Hachmann et al., ; Shin et al., ) including, enhanced peripheral (carotid body) chemoreflex sensitivity, which is a primary cause of sleep-disordered breathing in these subjects (Tester et al., ; Bascom et al., ). In contrast, studies in rats have found that mid-thoracic spinal cord injury is associated with enhanced cardiac sympathetic activity and cardiac sympathetic hyperinnervation that increases the susceptibility to life-threatening arrhythmias (Rodenbaugh et al., ; Collins et al., ; Lujan and DiCarlo, ; Lujan et al., , , , ). Studies in rats have also confirmed that hypoactivity of the CSC greatly enhances the likelihood of stroke in hypertensive models (Sadoshima et al., ; Sadoshima and Heistad, ; Werber and Heistad, ).

The effects of bilateral transection of the CSC (CSCX) or bilateral removal of the SCG (SCGX) have been investigated on a variety of physiological functions/variables in the mouse (Krieger et al., ; García et al., ; Kawaja and Crutcher, ; Pankevich et al., a,b; Karlsen et al., ; Lindborg et al., ; Ziegler et al., ). However, to our knowledge, no studies, in any species, have determined the roles of the CSC-SCG complex on ventilatory functions and responses to hypoxic challenges in vivo. The aim of this study was to determine the effects of sham-operated (SHAM) and bilateral CSCX (performed 4 days before testing) on resting ventilatory parameters in freely-moving adult male C57BL6 mice, and on their ventilatory responses to HXC using whole-body plethysmography as described previously (Palmer et al., a,b; Gaston et al., ; Getsy et al., ; Palmer et al., ).

The C57BL6 mouse has proven to be an invaluable murine model in which to investigate the physiological systems involved in ventilatory control processes (Tankersley et al., , ), and this strain is used widely to generate genetic knock-out mutants to investigate the molecular mechanisms underlying the responses of mice to hypoxic and/or hypercapnic gas challenges (Kline and Prabhakar, , Kline et al., ; Palmer et al., a). The ventilatory responses that will be described in the SHAM mice (during and following the HXC) were consistent with previous studies from our laboratory (Palmer et al., a,b, ; Gaston et al., ; Getsy et al., ). Our Getsy et al. () manuscript provides a detailed set of analyses of the ventilatory responses that occur in C57BL6, Swiss Webster and B6AF1 mice before, during, and after a brief HXC. We found that (1) the HVR in C57BL6 mice consists of an initial increase in frequency of breathing (Freq) followed by substantial decline (roll-off) toward pre-HXC values, whereas the Freq responses in Swiss Webster and B6AF1 mice were robust with minimal roll-off, and (2) the post-HXC (i.e., return to room-air) responses consisted of a rapid and sustained rise in Freq in C57BL6 mice, a sustained rise in the B6AF1 mice, which is known as a form of short-term potentiation (STP) (Powell et al., ), and a gradual return to pre-hypoxic challenge levels in the Swiss-Webster mice.

The ways and mechanisms by which activation of sympathetic nerves affects carotid body function under normoxic and hypoxic conditions have received considerable attention (Overholt and Prabhakar, ). For instance, compelling evidence shows that sympathetic nerve terminals innervate glomus cells and the microvasculature within the carotid bodies (Vázquez-Nin et al., ; McDonald, ; Verna et al., ), and that norepinephrine is the major neurotransmitter released by these terminals (Almaraz et al., ). The roles of norepinephrine and dopamine and α1-, α2-, and β-adrenoceptors and dopamine receptors have also been extensively studied. It is apparent that activation of sympathetic nerves can induce a multiplicity of effects within the carotid body. First, activation of the sympathetic nerves indirectly activate glomus cells by constricting arterioles (by activation α1-adrenoceptors and dopamine receptors) in the carotid body, effectively resulting in a hypoxic environment for glomus cells (Llados and Zapata, ; Majcherczyk et al., ; Matsumoto et al., ; Yokoyama et al., ). In addition, co-release of neuropeptide Y from sympathetic nerve terminals also reduces blood flow within the carotid bodies (Potter and McCloskey, ). In addition, activation of β-adrenoceptors and dopamine receptors on glomus cells directly activates these cells (Eldridge and Gill-Kumar, ; Lahiri et al., ; Gonsalves et al., ). On the other hand, intra-carotid artery infusions of norepinephrine depress resting chemoreceptor activity and also attenuate hypoxic excitation of the carotid body, effects mediated by α2-adrenoceptors (Kou et al., ; Pizarro et al., ; Prabhakar et al., ; Almaraz et al., ). Moreover, endogenous norepinephrine indirectly inhibits carotid body chemoafferent activity (Overholt and Prabhakar, ) via inhibition of glomus cell activity (presumably the release of excitatory neurotransmitters) by decreasing the magnitude and rate of macroscopic Ca2+ current influx (Overholt and Prabhakar, ). As such, it is likely to question how the absence of functional sympathetic input to the carotid bodies and other structures controlling ventilatory processes, would affect baseline parameters and the responses to HXC.

As mentioned, it has been established that HXC increases neural activity in the CSC and GGN (Lahiri et al., ; Matsumoto et al., , ). As such, it is proposed that HXC activates a carotid sinus (chemoafferent) nerve-brainstem-descending spinal cord pathway that increases pre-ganglionic sympathetic nerve activity, which in turn activates post-ganglionic neurons in the SCG, including those projecting to the carotid body via the GGN. It should be noted that there is compelling evidence that SCG cells are not directly responsive to hypoxia (Rigual et al., ; Nunes et al., ; Buckler and Turner, ; Gao et al., ; Bernardini et al., ), although there is equally compelling evidence that SCG cells [and in particular small intensely fluorescent (SIF) cells] are hypoxia-sensitive (Hanson et al., ; Brokaw and Hansen, ; Dinger et al., ; Strosznajder, ; Nunes et al., , ).

One principal finding of this study is that transection of the CSC is not equivalent to bilateral removal of the SCG (see Conclusion), suggesting multiple effects of hypoxia on neural signaling within the CSC-SCG pathway.

Materials and Methods

Permissions

All studies were carried out in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. ) revised in The protocols were approved by the Animal Care and Use Committee of Case Western Reserve University.

Animals

C57BL6 male mice were purchased from Jackson Laboratory (Bar Harbor, ME, United States). Mice were delivered pathogen free, and housed under specific-pathogen free conditions with a 12 h light-dark cycle. All procedures were performed in accordance with the National Institute of Health (NIH) guidelines for care and use of laboratory animals and were approved by the Institutional Animal Care and Use Committee at Case Western Reserve University.

Cervical Sympathetic Chain Transection (CSCX)

Adult mice (12 weeks) were anesthetized with an intraperitoneal injection of ketamine (80 mg/kg, Ketaset, Zoetis, Parsippany, NJ, mg/ml solvent) and xylazine (10 mg/kg, Akorn Animal Health, Lake Forest, IL, United States, 20 mg/ml solvent), and placed on a surgical station allowing body temperature to be maintained at 37°C via a heating pad (SurgiSuite, Kent Scientific Corporation, Torrington, CT, United States). The adequacy of anesthesia was regularly checked by nociceptive stimulus (e.g., a toe pinch). The SCG-CSC was identified behind the carotid artery bifurcation (Figure 1), and the CSC was cut using micro-scissors approximately 1 mm from the point where the CSC enters the SCG. In SHAM mice, the SCG-CSC was identified but not cut. The mice were allowed 4 days to recover from surgery. This time-point for recovery was chosen based on evidence that catecholamine levels in the carotid bodies, identified by tyrosine hydroxylase positive nerve terminals, are markedly reduced 3&#x;4 days after removal of the ipsilateral SCG (Mir et al., ; González-Guerrero et al., ; Ichikawa and Helke, ; Ichikawa, ). All mice were monitored for pain and distress every day following surgery. Mice were given an injection of the non-steroidal anti-inflammatory drug, carprofen (2 mg/kg, IP), 24 and 48 h post-surgery to reduce any pain or inflammation at the incision site. None of the mice showed any signs of pain or inflammation from the surgeries and began moving about the cages and eating and drinking approximately 1 h after surgery. Mice were weighed daily to ensure proper weight gain. We have determined that these injections of carprofen do not affect resting ventilation or the response to HXC on day 4 post-surgery (data not shown).

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Figure 1. Photograph of the left cervical sympathetic chain (CSC) and left superior cervical ganglion (SCG) of an adult C57BL6 mouse. The vagus nerve (Xth) and common carotid artery (CCA) are also shown. The scale bar is μm.

Whole-Body Plethysmography

On the day of study, mice were placed in individual whole-body plethysmographs (Buxco® Small Animal Whole Body Plethysmography, DSI a division of Harvard Biosciences, Inc., St. Paul, MN, United States) to continuously record ventilatory parameters in the freely-moving state, as described in detail previously (Palmer et al., a,b, ; Gaston et al., ; Getsy et al., ). The whole body plethysmography experiments were done by investigators (P.M.G, G.A.C, or Y-H.H) who were blinded to which surgery the mouse had undergone. The protocols were designed to ensure that the studies were done in a controlled fashion. More specifically, sub-groups of SHAM (n = 2) and CSCX (n = 2) mice were studied on the same day. The studies therefore took a total of 6 days over a 3 weeks period to study 11 SHAM and 12 CSCX mice. Because of the experimental paradigm, the collected data are likely to accurately define the effects of CSCX on resting ventilatory parameters and the response to HXC. The array of ventilatory parameters were chosen to provide an in-depth analysis of the differences in resting breathing patterns and responses to the hypoxic gas challenge in SHAM and CSCX mice (Solberg et al., ; Moore et al., ; Palmer et al., a,b, ; Gaston et al., ; Getsy et al., ; Villiere et al., ). Directly recorded parameters were (a) Freq, (b) tidal volume, (TV), (c) inspiratory time (Ti, duration of inspiration) and expiratory time (Te, duration of expiration), (d) end inspiratory pause (EIP, pause between end of inspiration and start of expiration) and end expiratory pause (EEP, pause between end of expiration and start of inspiration), (e) expiratory flow at 50% expired TV (EF50), (f) peak inspiratory flow (PIF) and peak expiratory flow (PEF), (g) relaxation time (decay of respiration to 36% of maximum PIF), (h) relative rate of achieving PEF (Rpef), and (i) rejection index (RI, % of non-eupneic breaths such as apneas per epoch). Calculated parameters were (a) minute ventilation (MV, Freq × TV), (b) Ti/Te and Ti/(Ti + Te), (c) PIF/PEF, (d) inspiratory drive (TV/Ti) and expiratory drive (TV/Te), (e) rejection index corrected for respiratory frequency (RI/Freq), and (f) the number of apneic pauses per epoch (Te/RT) With respect to the rejection index, the plethysmography system included a rejection algorithm that was set to reject breaths that did not reflect normal TV breathing, and as such rejected (a) abnormal breaths (abnormal balance of inspiratory and expiratory volumes), apneas, sighs, post-sighs, sniffs, and waveforms that most likely arose from activities such as grooming the face, hands, and rear (Getsy et al., ). Minimum TV was set at ml, minimum Ti was set at s, maximum Te was set at s, and the ratio of inspiratory volume/expiratory volume (i.e., volume balance) was set to a range of 90&#x;%. Values were rejected when they fell outside the above criteria and when (a) Ti was greater than 2 times Te, (b) PIF could not be distinguished from PEF, and (c) EF50, Rpef, or RT could not be computed (Getsy et al., ). All directly recorded parameters (i.e., Freq, TV, MV, Ti, Te, EIP, EEP, PIF, PEF, Rpef, relaxation time and rejection index) were extracted from the raw waveforms using proprietary Biosystem XA and FinePointe software (Data Sciences International, St. Paul, MN, United States) as described previously for mice (Palmer et al., a,b; Gaston et al., ; Getsy et al., ) and as detailed in the Data Sciences International/Buxco website reference to parameters provided by FinePointe Software using whole-body plethysmography. Data was extracted as individual data points (e.g., a Freq value for a particular 15 s epoch) and placed in excel spreadsheets.

Protocols for Hypoxic Gas Challenge

SHAM and CSCX mice were placed in the plethysmography chambers and allowed approximately 60 min to settle to allow for resting parameters to reach stable levels before the freely-moving mice were exposed to a 5 min HXC (10% O2 and 90% N2) and then re-exposed to room-air for 15 min.

Statistics

A data point before (15 min), during (5 min) and after (15 min) HXC was collected every 15 s. To determine the total responses (cumulative % changes from pre-HXC values) during HXC and return to room-air for each mouse, we summed the values recorded before and during the challenge and those upon return to room-air. Regarding the pre-HXC (baseline) phase, the breaths for each 15 s epoch were averaged over the last 5 min of the entire 15 min baseline recording period resulting in 20 values for each mouse that was averaged to give the resting value for each mouse. The mean and SEM for the 11 SHAM and 12 CSCX mice was then derived from the individual values. Similarly, twenty 15 s epoch values were derived during the 5 min hypoxic challenge and sixty 15 s epoch values were derived for the post-hypoxia (room-air) phase. Again, the mean and SEM for the 11 SHAM and 12 CSCX mice was derived from the individual values for the HXC and room-air phases under examination. We then determined the cumulative response for each mouse by the formulas, (a) total HXC response = (sum of the 20 values during HXC) &#x; (mean of the pre-HXC values × 20), and (b) total room-air response = (sum of the 60 values during room-air phase) &#x; (mean of the pre-HXC values × 60). We then determined the mean and SEM of the group data. We also calculated the total responses during the HXC 0&#x; sec epoch and &#x; sec epoch, in addition to the entire HXC 5 min (0&#x; sec epoch). All data are presented as mean ± SEM. All data were analyzed by one-way or two-way ANOVA followed by Student&#x;s modified t-test with Bonferroni corrections for multiple comparisons between means (Palmer et al., a,b).

Results

Resting Parameters

A summary of the mouse descriptors and resting ventilatory parameters is provided in Table 1. There were 11 mice in the SHAM group and 12 mice in the CSCX group. The ages and body weights of the two groups were similar to one another (P  , for both comparisons). Accordingly, no corrections for body weights were applied to the ventilatory data pertaining to volumes (e.g., TV and peak inspiratory and expiratory flows). There were no between-group differences for any recorded or calculated ventilatory parameter (P  , for all comparisons). In the following Figures 3&#x;10, the left panel of each row will show the actual values recorded before, during the 5 min hypoxic (10% O2 and 90% N2) challenge, and upon to room-air. The middle panels of each row will show the arithmetic change recorded over the first 60 s of hypoxic exposure. The right panels of each row will show the total response (% change from pre) during three epochs of the 5 min ( s) hypoxic challenge, namely 0&#x;, &#x;, and 0&#x; s. These epochs were those which best represented the dramatic differences between SHAM and SCGX mice (see Conclusion).

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Table 1. Baseline parameters in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX).

Hypoxic Challenges &#x; Frequency of Breathing, Tidal Volume and Minute Ventilation

Examples of respiratory waveforms during various stages of the experiment in a SHAM mouse and a CSCX mouse are shown in Figure 2. A time bar ( s) is shown in the bottom left of the figure. Resting Freq (pre-HXC values) was similar in both groups of mice. The initial HXC response (HXC at 15 s) was far greater in the CSCX mouse (&#x; breaths/min = breaths/min, +%) than the SHAM mouse (&#x; breaths/min = 48 breaths/min, +29%). The roll-off values at the end of the 5 min HXC were similar between the two groups (data not shown) as were the increases in Freq immediately upon return to room-air (RA at 15 s, + and +% for the SHAM and CSCX mouse, respectively) and at 5 min (RA at 5 min, + and +% for the SHAM and CSCX mouse, respectively).

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Figure 2. Example traces of respiratory waveforms during various stages of the experiment in a sham-operated (SHAM) mouse and in a mouse with bilateral cervical sympathetic chain transection (CSCX). A time bar ( s) is shown in the bottom left of the figure. HXC, hypoxic gas challenge. RA, room-air.

The changes in Freq, TV, and MV in SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure 3. As seen in the top row of panels, exposure to HXC in SHAM mice elicited a typical initial increase in Freq that was subject to pronounced roll-off (left panel). The responses in CSCX mice were essentially similar compared to SHAM except that the rise in Freq was significantly higher at the 15 s time-point (middle panel). The total increases in Freq in the three designated epochs (0&#x;, &#x;, and 0&#x; s) were similar in the SHAM and CSCX mice (right panel). The return to room-air elicited the expected dramatic increase in Freq in SHAM mice and CSCX mice (left panel), and the total room-air responses were similar in both groups (Table 2). As seen in the middle row of panels, exposure to the HXC elicited immediate and sustained increases in TV that were similar in most aspects in the SHAM and CSCX mice except that the total increase recorded during the &#x; s epoch were higher in CSCX than SHAM mice (right panel). The return to room-air elicited the expected small initial increase in TV followed by gradual decline toward baseline in SHAM and CSCX mice (left panel). The total room-air responses were similar in both groups (Table 2). As seen in the bottom row of panels, exposure to HXC in the SHAM mice elicited a typical initial increase in MV that was subject to a pronounced roll-off (left panel). The MV responses in CSCX mice were similar except that the rise in MV was significantly higher at the 15 s time-point (middle panel). The total increases in MV in the three designated epochs were similar in the SHAM and CSCX mice. The return to room-air elicited the expected dramatic increase in MV in SHAM and CSCX mice (left panel). The total room-air responses were similar in both groups (Table 2).

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Figure 3.(Left) Frequency of breathing, tidal volume and minute ventilation values before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

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Table 2. Total changes that occurred during the 15 min return to room-air.

Hypoxic Challenges &#x; Inspiratory Time and Expiratory Time

The changes in Ti and Te values in SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure 4. Exposure to HXC in SHAM mice elicited initial decreases in Ti and Te that were subject to roll-off (left panels). The decreases in Ti and Te occurred more rapidly in CSCX mice (middle panels), but the overall (total) responses for Ti and Te were similar in both groups (right panels). Return to room-air elicited a rapid transient decrease in Te, but a rapid and sustained decrease in Ti (left panels). The actual (left panels) and total room-air responses (Table 2) were similar in SHAM and CSCX mice. Additionally, the total Te responses upon return to room-air fell into two groups, those in which total Te fell and those in which Te rose (Table 2).

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Figure 4.(Left) Inspiratory time (Ti) and expiratory time (Te) values before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Hypoxic Challenges &#x; Inspiratory Time/Expiratory Time and Inspiratory Quotient

The changes in Ti/Te values and inspiratory quotients [Ti/(Ti + Te)] in SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure 5. The resulting changes in Ti and Te during HXC (Figure 4) translated into minor changes in Ti/Te values and inspiratory quotients in SHAM mice (left and middle panels) that were nevertheless significantly smaller in the CSCX mice (right panels). As seen in the left panels, the return to room-air resulted in substantial decreases in Ti/Te values and inspiratory quotients that gradually returned toward baseline values in both groups. As seen in Table 2, the changes in total Ti/Te and Ti/(Ti + Te) values upon return to room-air were similar in the SHAM and CSCX mice.

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Figure 5.(Left) Inspiratory time/expiratory time (Ti/Te) and inspiratory time/(inspiratory time + expiratory time) ratios [Ti/(Ti + Te)] before, during and after a 5 min hypoxic (HX, 10% O2 and 90 %N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Hypoxic Challenges &#x; End Inspiratory Pause, and End Expiratory Pause

The changes in EIP and EEP in SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure 6. The hypoxic challenge elicited a prompt decrease in EIP in SHAM and CSCX mice (top left panel). The initial (top middle) and total responses (top right panel) were similar in SHAM and CSCX mice. The HXC elicited a fall in EEP of about 2 min in duration in SHAM and CSCX mice (bottom left panel). Then EEP quickly returned to baseline values in the SHAM mice during the remainder of the hypoxic challenge, but went above baseline in the CSCX mice. The initial decrease in EEP during the HXC occurred faster in the CSCX mice (bottom middle panel) and the total changes in EEP (bottom right panel) reflected the biphasic changes described above. Return to room-air resulted in a gradual recovery of EIP toward baseline values, but substantial and variable increases in EEP (left panels) for both groups. As seen in Table 2, the total EIP and EEP responses upon return to room-air were similar in the SHAM and CSCX mice.

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Figure 6.(Left) End inspiratory pause (EIP) and end expiratory pause (EEP) values before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Hypoxic Challenges &#x; Inspiratory Drive and Expiratory Drive

The changes in inspiratory drive (TV/Ti) and expiratory drive (TV/Te) in the SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure 7. The HXC elicited prompt and sustained increases in both inspiratory and expiratory drives (left panels) that occurred more rapidly in CSCX mice (middle panels) although the total responses were similar in both groups (right panels). Return to room-air elicited initial increase in both inspiratory and expiratory drives in SHAM and CSCX mice that gradually returned toward baseline (left panels). The total room-air changes were similar in both groups (Table 2).

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Figure 7.(Left) Inspiratory Drive (TV/Ti) and Expiratory Drive (TV/Te) before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Hypoxic Challenges &#x; Peak Inspiratory and Expiratory Flows

The changes in PIF, PEF, and PIF/PEF ratios in the SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are shown in Figure 8. The HXC elicited prompt and sustained increases in PIF and PEF in SHAM and CSCX mice, with the PIF responses being of greater magnitude during the first half of the hypoxic challenge resulting in higher PIF/PEF ratios (left panels). The initial PIF responses during the HXC in the CSCX mice were similar to those in the SHAM mice, whereas the initial PEF responses were higher in CSCX mice, such that the expected increase in initial PIF/PEF ratios seen in SHAM mice did not occur in CSCX mice (middle panels). Total PIF responses to the HXC were similar in SHAM and CSCX mice, whereas the increases in PEF were higher in the CSCX mice compared to SHAM during each designated epoch (right panels). PIF/PEF ratios over the &#x; and 0&#x; s epochs were markedly lower in CSCX mice than in SHAM mice (bottom right panel). The return to room-air elicited initial increases in PIF and PEF in SHAM and CSCX mice, and the changes resulted in sustained increases in PIF/PEF ratios (left panels). Total PIF, PEF and PIF/PEF responses upon return to room-air were similar in both groups (Table 2).

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Figure 8.(Left) Peak Inspiratory flow (PIF) and Peak Expiratory Flow (PEF) values and PIF/PEF ratios before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Hypoxic Challenges &#x; EF50, Rpef, and Relaxation Time

The changes in EF50, Rpef, and RT values in the SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure 9. HXC elicited a robust increase in EF50 that occurred more rapidly in the CSCX mice, although the total responses were similar in the SHAM and CSCX mice (top row: left, middle, and right panels). HXC-induced initial brief increases in Rpef in SHAM and CSCX mice that were followed by sustained decreases (middle row: left panel). These Rpef responses occurred more rapidly in the SHAM mice than CSCX mice at the 45 sec time-point (middle row: middle panel), and the responses during the 0&#x; sec epoch were significantly smaller in the CSCX mice compared to SHAM, and significantly more reduced from baseline during the &#x; sec epoch and overall 0&#x; sec epoch (middle row: right panel). HXC elicited brief reductions in RT that occurred more rapidly in the CSCX mice than SHAM mice although the overall responses were similar in both groups (bottom row: left, middle, and right panels). The return to room-air elicited prompt increases in EF50 and Rpef that were associated with prompt decreases in RT (left panels). The overall room-air changes in EF50 were similar in both groups. The room-air changes in RT in the SHAM and CSCX mice fell into two categories of mice with roughly equal numbers, those in which RT was elevated and those in which RT was decreased. These two categories of changes were equivalent in the SHAM and CSCX mice. The changes in Rpef upon return to room-air also fell into two categories, roughly of equal numbers of mice, those in which Rpef was elevated and those in which Rpef was decreased. The values for those in which Rpef rose were significantly smaller in the CSCX mice than the SHAM mice (Table 2).

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Figure 9.(Left) EF50, Rpef and Relaxation Time values before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Hypoxic Challenges &#x; Rejection Index, Rejection Index/Frequency of Breathing, Apneic Pauses

The changes in RI, RI/Freq and Apneic Pause values in the SHAM and CSCX mice in response to the 5 min HXC and upon return to room-air are summarized in Figure The HXC elicited immediate, but short-lived increases in RI and RI/Freq values that were not accompanied by increases in the numbers of apneic pauses, which on the contrary, showed relatively transient decreases in the earlier stage of the HXC (left panels). The increases in RI and RI/Freq occurred more rapidly in the CSCX mice, but the overall changes were similar in the SHAM and CSCX mice (top and middle rows: middle and right panels). The changes in apneic pauses during the HXC were similar in the SHAM and CSCX mice (bottom row). The return to room-air was associated with rapid and substantial increases in RI, RI/Freq and apneic pauses (left panel) that were similar in magnitude in the SHAM and CSCX mice (Table 2).

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Figure (Left) Rejection Index (RI, Rinx) values, RI/frequency ratios and Apneic Pauses (AP) before, during and after a 5 min hypoxic (HX, 10% O2, 90% N2) gas challenge in sham-operated (SHAM) mice and in mice with bilateral transection of the cervical sympathetic chain (CSCX). (Middle) Responses (expressed as % of pre-values) during the first minute of HX challenge in SHAM and CSCX mice. (Right) Total responses (sum of all % changes from pre) during the first s, between &#x; s and 0&#x; s. Data are expressed as mean ± SEM. P , significant response. &#x;P , CSCX versus SHAM. There were 11 mice in the SHAM group and 12 mice in the CSCX group.

Discussion

Due to the multiplicity and often opposing effects of sympathetic nerves/catecholamines in the carotid bodies (see Introduction), the question as to how the absence of functional sympathetic input to the carotid bodies and other structures controlling ventilatory processes would affect baseline parameters and the responses to HXC was not easy to predict. The major finding that many of the ventilatory responses (e.g., increases Freq, PEF and expiratory drive) during HXC occurred faster in CSCX mice suggests that activation of SCG sympathetic input to the carotid bodies of these mice normally blunts the initiation of these responses. Accordingly, it appears that loss of sympathetic input/catecholamine-induced suppression of carotid body activity (Kou et al., ; Pizarro et al., ; Prabhakar et al., ; Almaraz et al., ; Overholt and Prabhakar, ) out ways the loss of mechanisms which promote hypoxic responses including vasoconstriction in arterioles associated with glomus cells (Llados and Zapata, ; Eldridge and Gill-Kumar, ; Majcherczyk et al., ; Lahiri et al., ; Matsumoto et al., ; Gonsalves et al., ; Potter and McCloskey, ; Yokoyama et al., ). Deeper mechanistic insights would certainly come from studies designed to evaluate blood flow responses in the mouse carotid body and cerebral circulation in SHAM and CSCX mice during HXC and how systemic hemodynamic responses in these mice may influence the expression of the ventilatory responses.

Ventilatory Responses During Hypoxic Gas Challenge

This study demonstrates that HXC in adult male C57BL6 mice elicits a complex array of ventilatory responses over and above what have been reported previously. In response to the HXC, the SHAM C57BL6 mice displayed an increase in Freq that was subject to roll-off and which was accompanied with decreases in Ti, Te, EEP, Rpef and RT that were also subject to roll-off. In contrast, the decrease in EIP was sustained throughout HXC. HXC was also associated with robust and sustained increases in TV, MV, PIF, PEF, EF50, and inspiratory and expiratory drives. The question arises as to why some ventilatory parameters in C57BL6 mice are subject to roll-off whereas others are not. First, it should be noted that the C57BL6 mouse is widely considered to be a normal healthy model to study the physiology of cardiorespiratory systems, and is used to generate genetically-engineered mice to study the mechanisms involved in cardiorespiratory-thermoregulatory control processes, including responses to HXCs (Tankersley et al., ; Campen et al., , ; Palmer et al., a,b; Tewari et al., ; Gaston et al., ). Despite considerable normal physiology, the C57BL6 mouse is of major interest to sleep-apnea researchers because it displays disordered breathing (irregular breathing patterns, including apneas and sighs) and cardiovascular disturbances during sleep and wakefulness, and shows disordered breathing upon return to room-air after exposure to HXC (Han and Strohl, ; Han et al., , ; Tagaito et al., ; Yamauchi et al., a,b,c, ; Getsy et al., ). The genetic (Tankersley et al., , , ; Tankersley, , ; Han et al., , ; Tagaito et al., ; Yamauchi et al., b) and neurochemical processes (Tankersley et al., ; Price et al., ; Groeben et al., ; Yamauchi et al., a,c, ; Moore et al., , ), underlying the breathing patterns of C57BL6 mice, and the responses to HXC are well studied. The potential role of structural differences in the carotid bodies has also been studied (Yamaguchi et al., , ; Chai et al., ). Nonetheless, evidence that the breathing patterns of C57BL6 mice and their responses to HXC have a strong genetic component has not posed an explanation as to why some ventilatory components, such as ventilatory timing (e.g., Freq and EEP) and mechanics (e.g., Rpef and RT) are subject to roll-off whereas other timing (e.g., EIP) and mechanics (e.g., PIF, PEF, and EF50) are not. Regardless of the explanation, understanding the importance of each of these ventilatory responses to HXC will help us better understand the processes by which breathing disorders occur in disease states and point to therapeutic strategies.

Resting ventilatory parameters (19 directly recorded or calculated variables) were similar in the SHAM and CSCX mice. This would suggest that (presumed) loss of input from the SCG to structures controlling breathing, such as the carotid bodies, upper airway and brainstem structures (see Introduction), do not obviously effect ventilatory timing or mechanics or the quality of breathing (e.g., occurrence of non-eupneic breathing events, such as apneic pauses) in C57BL6 mice. Again, the caveat is that these studies were performed only 4 days after CSCX, and it would seem possible that changes in baseline ventilatory performance would occur at longer post-CSCX time-points. Nevertheless, there were numerous important differences in the responses of CSCX and SHAM mice to the HXC. For example, the increases in Freq (and associated decreases in Ti, Te, EEP, and RT) and the increases in MV, inspiratory and expiratory drives, PEF and EF50 occurred more quickly in CSCX mice than in SHAM mice. Although both PIF and PEF increased during HXC in the CSCX mice, the PIF/PEF ratio fell because the rise in PEF exceeded that of the rise in PIF, and the PIF/PEF ratio fell more dramatically, and to a greater extent, in the CSCX mice because of the exaggerated rise in PEF in the CSCX mice. The overall (total) responses in the SHAM and CSCX mice to the HXC were similar to one another with some important exceptions. Specifically, the overall increases in TV during the latter half of the hypoxic challenge appeared greater in the CSCX mice, and as mentioned above, the PIF/PEF ratio fell and to a greater extent in the CSCX mice. Moreover, the total decreases in Ti/Te and respiratory quotient [Ti/(Ti + Te)] observed in the SHAM mice were absent in CSCX mice. These findings clearly demonstrate that CSC-SCG input to respiratory control structures influence the ventilatory responses to HXC in C57BL6 mice.

Ventilatory Responses Upon Return to Room-Air

Following exposure to HXC, the return to room-air resulted in respiratory patterns that can be classified as short-term potentiation, in which ventilation remains elevated (Powell et al., ; Getsy et al., ) or post-hypoxic frequency decline, in which breathing frequency falls below baseline (Dick and Coles, ). Our C57BL6 mice displayed robust short-term potentiation upon return to room-air, accompanied by a substantial prolonged phase of disordered breathing (e.g., elevated rejection index). The mechanisms responsible for post-HXC disordered breathing have received considerable investigation, and at present, evidence is in favor of disturbances in central signaling (Wilkinson et al., ; Strohl, ) including, the pons area of the brainstem (Coles and Dick, ; Dick and Coles, ) rather than processes within the carotid bodies (Vizek et al., ; Brown et al., ), even though it is evident that carotid body chemoafferents play a vital role in the expression of disordered breathing such as, sleep apnea (Smith et al., ). The post-HXC responses were similar in our SHAM and CSCX mice, suggesting that diminished input to the SCG and (presumably decreased activity of SCG cells) do not have an obvious impact on the post-HXC responses, including the disordered breathing. The one exception was that total Rpef responses (positive rather than negative responders) (Table 2) after return to room-air were smaller in the CSCX mice than in the SHAM mice. This suggests that CSC-SCG activity is normally a positive factor in achieving maximal Rpef upon recovery from a HXC challenge in C57BL6 mice. The major differences with respect to the initial (i.e., first 60 sec of the HXC) changes in Freq, MV, EEP, rejection index (Rinx), and Rinx/Freq did occur during the first 15 s although other differences between the SHAM and CSCX groups occurred at 30 s for relaxation time and inspiratory drive; 45 s for Rpef; 15 and 30 s for Ti and Te; 30, 45, and 60 s for PEF, PEF/PIF and EF50; and 15, 30, 45, and 60 s for expiratory drive.

Study Limitations

A limitation of this study is that it only provides a snap-shot of the temporal changes in ventilatory responses to HXC after bilateral CSCX. Therefore, it is imperative to study what the patterns of ventilatory responses to HXC would be at longer time-points post-CSCX. We chose to test the mice 4 days after CSCX surgery in order to compare with our findings related to the effects of bilateral removal of the SCG on HXC (see Conclusion). The 4-day recovery period was chosen for our other study as it would be the earliest time in which SCGX would result in virtually complete loss of sympathetic terminals within the carotid bodies (Mir et al., ; González-Guerrero et al., ; Ichikawa and Helke, ; Ichikawa, ). We have yet to establish whether this loss is true in mice, and also whether there is significant loss of sympathetic terminals in other structures, such as the brainstem. This presence/absence of intact terminals is vital since it is established that hypoxia elicits action potential/extracellular Ca2+-independent release of catecholamines from sympathetic nerve terminals (Schömig et al., ; Chahine et al., ; Kurz et al., , ; Du et al., ). This evidence is especially relevant to our study since CSCX would presumably diminish activity of post-ganglionic neurons within the SCG without causing the degeneration of these post-ganglionic nerve terminals. Moreover, substantial data has shown that spinal cord damage, which markedly reduces pre-ganglionic outflow, does not necessarily eliminate all post-ganglionic sympathetic nerve activity (Meckler and Weaver, ; Qu et al., ; Stein and Weaver, ; McLachlan, ). Nonetheless, to our knowledge this has not been studied for T1&#x;T4 spinal cord-CSC-SCG pathways. It would seem reasonable to suggest that since we transected the left and right CSC and therefore the pre-ganglionic fibers in these nerves, we could expect that (1) the activity of a post-ganglionic fibers emanating from the SCG including those to the carotid bodies would be markedly if not totally diminished, and (2) HXC would not be able to elicit centrally-mediated changes in SCG neuronal activity. However, it certainly remains possible that HXC directly activates post-ganglionic neurons denervated of pre-ganglionic input and/or more likely, direct release of neurotransmitters from sympathetic nerve terminals themselves as demonstrated in the heart and saphenous veins (Dart and Riemersma, ; Kamath et al., ; Chahine et al., ; Santos et al., ), if vesicular release mechanisms are intact.

Conclusion

Our data show that under baseline (normoxic) environmental conditions, the potential loss of active sympathetic input to the carotid bodies (via CSCX-induced quiescence of post-ganglionic projections to the carotid body) does not have a noticeable effect on ventilatory parameters, which suggests that resting activity (e.g., neurotransmitter release) of carotid body glomus cells is not altered in a way that would lead to activation of carotid body chemoafferents and therefore enhancement of breathing. In contrast, our data shows that CSCX does alter the initial hypoxic ventilatory response and therefore suggests that diminished SCG input to the carotid bodies (or other targets, such as those in the brainstem) does influence the ability of glomus cells and/or other neuronal structures to respond to HXC. Planned studies involving bilateral transection of ganglioglomerular nerves (that project only to the carotid bodies and carotid sinus) will help to establish the relevant neuronal pathways/mechanisms. Overall, this novel data suggest that the CSC may normally provide inhibitory input to peripheral (e.g., carotid bodies) and central (e.g., brainstem) structures that are involved in the ventilatory responses to HXC in C57BL6 mice. Moreover, the results of our CSCX study lend support to the concept that a loss of CSC-SCG activity may be involved in the etiology of ventilatory disorders, such as sleep-disordered breathing. With respect to mechanistic insights provided by our data, it would be reasonable to assume that post-ganglionic sympathetic nerves innervating the ipsilateral carotid bodies (and other targets such as those within the brain) would be quiescent following CSCX as a result of the loss of pre-ganglionic input to the SCG. Whether the presumed diminution of sympathetic activity alters the expression of functional proteins (i.e., tyrosine hydroxylase, catecholamine-containing vesicles, and fusion proteins mediating vesicular exocytosis) within the sympathetic nerve terminals themselves and/or the target tissues, such as glomus cells in the carotid bodies needs to be addressed in future experiments. This is especially important since these post-transection adaptations in protein expression, while not being noticeable at rest (i.e., under normoxia) may have a direct effect on the ability of glomus cells (for example) to respond to the hypoxic challenge and/or secrete neurotransmitters. We have data that demonstrates that removal of the SCG has dramatically augmented effects on HXC compared to CSCX (unpublished observations). This raises important questions as to whether HXC may directly alter the activity of SCG neurons independently of the CSC input. One key question pertains to which of the SCG projections normally driven by the CSC are responsible for mediating the neuromodulatory effects of the CSC on the processes that drive ventilatory responses to HXC. As detailed in the Introduction section, the direct projections of the SCG to structures that control ventilation are extensive and include projections to the carotid bodies via the GGN. In order to better determine which of the post-ganglionic projections of the SCG regulate the ventilatory responses to HXC, we are currently planning to perform studies in mice in which the major post-ganglionic branches of the SCG, namely the internal carotid nerves, external carotid nerves and GGN are transected. We intend to perform these studies 4, 14, and 30 days post-transection in C57BL6 mice and in other strains, such as Swiss-Webster and A/J mice (Getsy et al., ) to determine temporal and genetic aspects of the role of the CSC-SCG complex in the control of ventilation and the responses to HXC.

The data in this study demonstrates that the primary effect of CSCX appears to be changes in the immediate responsiveness to the HXC. For example, the increases in Freq (and associated decreases in Ti, Te, and EEP), MV, expiratory drive, and rejection index occurred more rapidly in CSCX mice than SHAM mice (changes at 15 sec were significant in CSCX mice but not SHAM mice and between-group differences for expiratory drive were maintained at 15, 30, 45, and 60 s). In contrast, between-group differences in the responses of relaxation time, Rpef, PIF, PEF, PEF/PIF, EF50, and inspiratory drive, were evident at 30, 45, or 60 s with between-group differences for PEF, PEF/PIF, and EF50 being evident at 30, 45, and 60 s. Additionally, it is important to remember for the interpretation of the effects of CSCX that it was evident that the initial increases in TV and PIF during HXC were similar in SHAM and CSCX mice. Taken together, it is apparent that the loss of post-ganglionic SCG input to structures, such as the carotid body and brainstem, has a strong impact on ventilatory performance in C57BL6 mice. The findings that the decreases in Ti and Te were larger in the CSCX mice than the SHAM mice suggests that SCG input to neural structures regulating ventilatory timing events equally affect inspiratory and expiratory control processes. However, with respect to flow parameters it was evident that (a) the increases in TV were similar in SHAM and CSCX mice, (b) the responses of PIF and inspiratory drive in CSCX mice were minimally different from SHAM mice, whereas the changes in PEF, EF50, PEF/PIF and expiratory drive were substantially different between the groups. As such, CSCX appears to have much more of an influence on PEF than PIF. Whether this pattern of effects is due primarily to altered carotid body function must await more definitive studies in which, for example, the effects of bilateral GGN transection are investigated.

The data in the present manuscript and in that of our recent study which investigated the effects of bilateral SCGX (unpublished observations) provides the beginning of understanding how the loss of pre-ganglionic and/or post-ganglionic fibers in the CSC-SCG complex affect resting ventilatory parameters and the responses to HXC. On-going studies will extend our investigations by (1) determining how expression of proteins in sympathetic nerve terminals (e.g., tyrosine hydroxylase, fusion proteins, and vesicular stores of norepinephrine) and carotid body glomus cells (e.g., tyrosine hydroxylase, voltage-gated Na+, K+, and Ca2+-channels) change after CSCX, and (2) differentiating the effects of sympathetic input to various areas by performing ventilatory studies in mice with (a) transection of the left and right internal carotid nerves (a major post-ganglionic SCG trunk), (b) transection of the left and right external carotid nerves (the other major post-ganglionic SCG trunk), and (c) transection of the left and right ganglioglomerular nerves (a branch of the external carotid nerve), which projects only to the carotid bodies and carotid sinus.

Data Availability Statement

The raw data supporting the conclusions of this article will be made available by the authors, without undue reservation.

Ethics Statement

The animal study was reviewed and approved by Case Western Reserve University Institution&#x;s Animal Care and Use Committee.

Author Contributions

PG, GC, Y-HH, and SL conceived and designed the study. PG performed the mouse surgeries. PG and GC performed the plethysmography studies. PG and SL analyzed the data and prepared the figures. All authors contributed to writing the manuscript, and revised, read, and approved the final version of the manuscript.

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Funding

This study was funded by an NIH-SPARC award to SL (10T20D; Functional Mapping of the afferent and Efferent Projections of the Superior Cervical Ganglion Interactome).

Acknowledgments

The authors wish to thank Dr. James N. Bates (Department of Anesthesia, University of Iowa) for his critical comments about the manuscript and helping with the clinical perspectives of the study.

Abbreviations

CSC, cervical sympathetic chain; CSCX, cervical sympathetic chain transection; SCG, superior cervical ganglion; SCGX, superior cervical ganglionectomy; HXC, hypoxic gas challenge; Freq, frequency of breathing; TV, tidal volume; MV, minute ventilation; Ti, inspiratory time; Te, expiratory time; EIP, end inspiratory pause; EEP, end expiratory pause; PIF, peak inspiratory flow; PEF, peak expiratory flow; RT, relaxation time; EF50, expiratory flow at 50% expired total volume; Rpef, rate of achieving peak expiratory flow.

Footnotes

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Sours: https://www.frontiersin.org/articles//fphys/full

Ganglion superior cervical

Block of the Superior Cervical Ganglion, Description of a Novel Ultrasound-Guided Technique in Human Cadavers

Abstract

Objective.

Injection of opioids to the superior cervical ganglion (SCG) has been reported to provide pain relief in patients suffering from different kinds of neuropathic facial pain conditions, such as trigeminal neuralgia, postherpetic neuralgia, and atypical facial pain. The classic approach to the SCG is a transoral technique using a so-called “stopper” to prevent accidental carotid artery puncture. The main disadvantage of this technique is that the needle tip is positioned distant from the actual target, possibly impeding successful block of the SCG. A further limitation is that injection of local anesthetics due to potential carotid artery puncture is contraindicated. We hypothesized that the SCG can be identified and blocked using ultrasound imaging, potentially increasing precision of this technique.

Interventions.

In this pilot study, 20 US-guided simulated blocks of the SCG were performed in 10 human cadavers in order to determine the accuracy of this novel block technique. After injection of mL of dye, the cadavers were dissected to evaluate the needle position and coloring of the SCG.

Results.

Nineteen of the 20 needle tips were located in or next to the SCG. This corresponded to a simulated block success rate of 95% (95% confidence interval 85–%). In 17 cases, the SCG was completely colored, and in two cases, the caudal half of the SCG was colored with dye.

Conclusions.

The anatomical dissections confirmed that our ultrasound-guided approach to the SCG is accurate. Ultrasound could become an attractive alternative to the “blind” transoral technique of SCG blocks.

Interventional, Neuropathic Pain, Pain Management, Sympathetic Block, Ultrasound

Introduction

The superior cervical ganglion (SCG) is the most cranial part of the sympathetic chain and provides sympathetic innervation to the face and head . It is the largest of the cervical ganglia, fusiform in shape, often flattened, and located in a plication of the prevertebral (deep cervical) fascia anterior to the longus capitis muscle and dorsal to the internal carotid artery. At the C3 level, it is located posteromedial to the vagus nerve. In contrast with the other two cervical ganglia, it is absolutely constant. It may reach caudally as far as to the upper border of the fourth cervical vertebra, but the main portion is found at the level of the transverse processes of the second and third cervical vertebrae .

Injection of low-dose buprenorphine to different sympathetic ganglia has been termed “GLOA” (ganglionic local opioid analgesia) and is used to treat various chronic pain states. Even though clear evidence of a specific therapeutic effect of buprenorphine injected next to the SCG is lacking , the procedure has been reported to provide pain relief without side effects in patients suffering from different kinds of neuropathic facial pain conditions, such as trigeminal neuralgia, postherpetic neuralgia, and atypical facial pain . The standard technique described to block the SCG is a blind, transoral approach where a needle is inserted at a slightly retrotonsillar location through the dorsolateral pharyngeal wall using a so-called “stopper,” preventing the needle from penetrating the pharyngeal wall deeper than 1 cm and avoiding accidental carotid artery puncture (Figure ). Using this method, the area of the SCG is targeted at C2 and/or intersection C1/2 level. The potential risk of an unnoticed carotid artery puncture (not least due to a possible tortuous course/coiling of the ICA) remains the main reason why the application of local anesthetics with this blind, transoral approach is contraindicated .

Figure 1

Scheme of the needle path of the blind transoral approach (at level of C2) using a stopper (1) as compared with our novel ultrasound-guided posterolateral approach (2) to the superior cervical ganglion (at C3 level). ICA = internal carotid artery; IJV = internal jugular vein; X = vagus nerve; SCG = superior cervical ganglion; LCM = longus capitis muscle; NR = nerve root C2/C3; TP = transverse process of C2/C3; VB = vertebral body of C2/C3.

Figure 1

Scheme of the needle path of the blind transoral approach (at level of C2) using a stopper (1) as compared with our novel ultrasound-guided posterolateral approach (2) to the superior cervical ganglion (at C3 level). ICA = internal carotid artery; IJV = internal jugular vein; X = vagus nerve; SCG = superior cervical ganglion; LCM = longus capitis muscle; NR = nerve root C2/C3; TP = transverse process of C2/C3; VB = vertebral body of C2/C3.

In addition to the blind approach, a fluoroscopically guided technique to block the SCG has been described as well. The fluoroscopic technique is performed in a similar manner as the paratracheal approach to the stellate ganglion, although at a more cranial paratracheal location at the level of the transition of the vertebral body of C3 to its transverse process . Treggiari and coworkers blocked the SCG with bupivacaine and clonidine using this fluoroscopic-guided approach in order to relief cerebral vasospasm following subarachnoidal bleeding . All the patients displayed improved cerebral perfusion in the confirmatory angiography. Besides the issue of radiation exposure of patient and staff, the main disadvantage of fluoroscopic imaging is that it does not allow visualization of potentially hazardous blood vessels in the needle path.

Ultrasound imaging offers new opportunities for superior visualization of this large neural structure. Due to its large size, the SCG is expected to be visualized with ultrasound, and hence, it could be blocked under ultrasound guidance. To our knowledge, based on PubMed and Internet searches, this has not been described.

We performed a feasibility study in 10 human cadavers in order to determine the ability and accuracy to visualize and “block” the SCG with ultrasound imaging. After identification of the target structure with ultrasound imaging, we injected a small amount of dye and verified the selective coloring of the SCG by anatomical dissection.

Materials and Methods

The cadavers had been embalmed in glycerol and alcohol, as previously described , and were in legal custody of the Department of Anatomy, Histology, and Embryology (Innsbruck, Austria). Institutional approval for the procedure was obtained. An Esaote MyLab® ultrasound system (Esaote Biomedica, Cologne, Germany) with a mm microconvex array transducer with a maximal resolution of 8 MHz was used.

The cadavers were positioned supine, and the head rotated in a slightly contralateral direction. First, with the transducer held in a transverse plane, the transverse process of C6 was located by identifying its prominent anterior tubercle (Chassaignac's tubercle). Thereafter, the probe was carefully moved in a cephalad direction, and each transverse process was counted until the level C3 was identified. Here, the transducer was moved medially until the landmarks of the longus capitis muscle and internal carotid artery were identified. Between these two structures, we searched for a slightly hypoechoic oval to round (in the transversal plane) and fusiform structure (in the consecutively performed longitudinal scan), which we hypothesized to be the SCG. After measuring its maximal visible length and anteroposterior diameter, a G needle (SonoTAP needle 80 mm, Pajunk Medical Products, Geisingen, Germany) was introduced from a posterolateral approach with an in-plane technique using real-time ultrasound guidance. The needle was directed into or immediately adjacent to the structure appearing to be the SCG.

Once the needle was in position, mL of indocyanine green (ICG) (ICG Pulsion, Pulsion Medical Systems AG, Munich, Germany) was injected. Immediately after injection, the cadavers were carefully dissected by the independent anatomist (B.M.) under manual fixation of the needle to show the actual position of the needle tip and the spread of the dye. The needle position was defined as correct if its tip was in contact to or located within the targeted ganglion and the SCG itself was colored.

Results

The characteristics of the 10 examined cadavers were: sex: four female, six male; median age at death 80 years (range 66–91); median body mass index 23 (range –27) kg/m2.

We targeted both sides in each of the 10 cadavers; hence, 20 simulated SCG blocks were performed. Anatomical dissection revealed that in 19 cases, the needle tip was located in or next to the SCG; thus, the success rate of our simulated SCG block was 95% (95% confidence interval 85–%). Of the 19 successfully reached SCGs, 17 were completely colored by the dye, and in two, the caudal half of the ganglion was colored.

In the single case, where the SCG was missed, the needle tip was identified in a too caudal position at C4 and located in between the internal and external carotid artery.

In the 19 correctly identified SCGs, the median of the maximal anteroposterior diameter as measured with ultrasound was 2 mm (range 1–4), and the median measurable length was 10 mm (range 7–17).

Figure shows an example of a successful simulated SCG block as confirmed by anatomical dissection, and Figure shows the corresponding ultrasound image.

Figure 2

An example of the injection site after anatomical dissection. The dye was injected into the superior cervical ganglion (SCG), which has been colored completely. Note that the internal carotid artery (ICA) has been shifted anteriorly in order to enable visualization of the SCG. LCM = longus capitis muscle.

Figure 2

An example of the injection site after anatomical dissection. The dye was injected into the superior cervical ganglion (SCG), which has been colored completely. Note that the internal carotid artery (ICA) has been shifted anteriorly in order to enable visualization of the SCG. LCM = longus capitis muscle.

Figure 3

Ultrasound image and measurement (+−−−−+) of the SCG in a cadaver. Both the superior cervical ganglion (SCG) and the internal carotid artery (ICA) have been cut in a longitudinal plane. Note the typical hypoechoic appearance and fusiform shape of the SCG. The lumen of the ICA is partially compressed; this is a common observation when scanning these specially embalmed cadavers.

Figure 3

Ultrasound image and measurement (+−−−−+) of the SCG in a cadaver. Both the superior cervical ganglion (SCG) and the internal carotid artery (ICA) have been cut in a longitudinal plane. Note the typical hypoechoic appearance and fusiform shape of the SCG. The lumen of the ICA is partially compressed; this is a common observation when scanning these specially embalmed cadavers.

Discussion

This investigation is the first description of an ultrasound-guided approach to the SCG. In the 10 examined cadavers, the new technique proved to be highly accurate with a simulated block success rate of 95%.

The main advantages of this novel approach is that the needle tip can be positioned under ultrasound guidance directly next to the SCG, as opposed to the classic transoral technique where the needle tip is located anterior to the internal carotid artery and hence distant from the SCG.

A previous study by Feigl and coworkers investigated the simulated success rate of the classic transoral approach to the SCG . They were able to demonstrate that volumes as little as 1–2 mL spread sufficiently to the SCG, but a strictly lateral puncture direction was considered as crucial because only a slightly too medial puncture direction resulted in penetration of the prevertebral fascia with resulting block failure.

Other potential advantages of this ultrasound-guided approach are as follows: Application of local anesthetics to the SCG could be reconsidered with this novel approach, as accidental internal carotid artery puncture (due to a tortuous course and/or coiling of the artery ) should be prevented with ultrasound imaging.

As opposed to the fluoroscopic approach described in the study of Treggiari and coworkers , where block of the SCG was performed in order to alleviate cerebral vasospasm after subarachnoid hemorrhage (SAB), the most important advantage of ultrasound guidance is that it avoids radiation exposure and can be performed outside lead-shielded facilities. Even though radiation exposure of a single SCG block may be low, usually several blocks on different occasions are required to relief facial pain or bring other acceptable results, which may lead to considerable radiation exposure of patients and staff . A further advantage to fluoroscopy (or the blind transoral approach) is that ultrasound imaging enables identification of other potentially hazardous structures in the needle path (besides the already-mentioned internal carotid artery), such as other relevant arteries (e.g., ascending pharyngeal artery) and nerves (e.g., the vagus and the accessory nerves). Even though these structures are visible using ultrasound, needle tip position should be as close as possible to the mentioned prevertebral muscles to avoid potential damage.

Some technical considerations need to be addressed. Standard high-resolution linear ultrasound probes are not suited to identify the SCG using our technique because of the difficulty of trying to scan the cranial cervical spine in a transverse plane in the presence of the mandible. A transversal scan is mandatory to introduce the needle using an in-plane technique in order to prevent penetration of the internal carotid artery. Hence, we used a considerably smaller mm microconvex array transducer to overcome this issue. Despite the lower resolution, the large SCG could be identified in most cases. However, its actual length, as assessed by a longitudinal scan, could not reliably be measured in most cases because of the difficulty of trying to scan the cranial cervical spine in the presence of the mandible impairing visibility of the cranial ending of the SCG. This explains the considerably smaller lengths we assessed by ultrasound measurements (median 10 mm, range 7–17 mm) as opposed to values described in the anatomical literature (25–30 mm ).

We conclude that in our pilot study performed on 10 human cadavers, the SCG could reliably be identified and a needle placed in or immediately next to it using a novel ultrasound-guided approach. Further clinical research is required to compare this technique to previous techniques and to reconsider the injection of local anesthetics. This presumably more reliable ultrasound-guided approach could also serve as platform for the planning of better future clinical outcome studies.

References

© American Academy of Pain Medicine

Sours: https://academic.oup.com/painmedicine/article/14/5//
Neurology - Sympathetic Nervous System

Sympathetic Innervation to the Head and Neck

The sympathetic nervous system is a division of the autonomic nervous system. It is involuntary, and acts with the parasympathetic system to maintain body homeostasis.

The actions of the sympathetic nervous system are associated with the 'fight or flight' response.

In this article, we shall look at the anatomy of the sympathetic innervation to the head and neck - its structure, anatomical course, and its clinical correlations.


Anatomical Structure and Course

The sympathetic fibres to the head and neck begin in the spinal cord. They originate from the thoracic region (T), and therefore need to ascend to reach the structures in the head and neck.

After leaving the spinal cord, the fibres enter the sympathetic chain. This structure spans from the base of the skull to the coccyx, and is formed by nerve fibres and ganglia (collections of nerve cell bodies). There are three ganglia within this chain that are of interest - the superior, middle and inferior cervical ganglia. The sympathetic fibres synapse with these ganglia, with post ganglionic branches continuing into the head and neck.

Each of the three ganglia are related to specific arteries in the head and neck. The post-ganglionic fibres hitch-hike along these arteries (and their branches) in order to reach their target organs.

We shall now look at the structure and function of the ganglia in more detail.

Note: In some individuals, the middle cervical ganglion is often absent and the inferior cervical ganglion is often fused with the first thoracic ganglion, as a result is known as the cervicothoracic ganglion. In addition to this, the superior and middle cervical ganglia are commonly connected together.

Superior Cervical Ganglion

[caption id="attachment_" align="alignright" width=""]Fig - The superior, middle and inferior cervical gangliaFig - The superior, middle and inferior cervical ganglia[/caption]

The superior cervical ganglion is located posteriorly to the carotid artery, and anterior to the C vertebrae. Several important post-ganglionic nerves originate from here:

  • Internal carotid nerve - hitch-hikes along the internal carotid artery, forming a network of nerves. Branches from the internal carotid plexus innervate structures in the eye, the pterygopalatine artery and the internal carotid artery itself.
  • External carotid nerve - hitch-hikes along the common and external carotid arteries, forming a network of nerves. It innervates the smooth muscle of the arteries.
  • Nerve to pharyngeal plexus - combines with branches from the vagus and glossopharyngeal nerves to form the pharyngeal plexus.
  • Superior cardiac branch - contributes to the cardiac plexus in the thorax.
  • Nerves to cranial nerves II, III IV, VI and IX.
  • Gray rami communicantes - distributes sympathetic fibres to the anterior rami of C1-C4.

Middle Cervical Ganglion

The middle cervical ganglion is absent in some individuals. When present, it is located anteriorly to the inferior thyroid artery and the C6 vertebra. Its postganglionic fibres are:

  • Gray rami communicantes - distributes sympathetic fibres to the anterior rami of C5 and C6.
  • Thyroid branches - travel along the inferior thyroid artery, distributing fibres to the larynx, trachea, pharynx and upper oesophagus.
  • Middle cardiac branch - contributes to the cardiac plexus in the thorax.

Inferior Cervical Ganglion

The inferior cervical ganglion is situated anteriorly to the C7 vertebra. It is occasionally fused with the first thoracic vertebrae, forming the cervicothoracic ganglion. There are three post-ganglionic fibres that arise from this ganglion:

  • Gray rami communicantes - distributes sympathetic fibres to the anterior rami of C7, C8 and T1.
  • Branches to the subclavian and vertebral arteries - Theseinnervate the smooth muscle present in the arteries.
  • Inferior cardiac nerve - contributes to the cardiac plexus in the thorax.

[start-clinical]

Clinical Relevance: Horner's Syndrome

The sympathetic fibres can be stretched or damaged along their course to the head and neck. If these nerves are unilaterally disturbed, it produces a triad of main symptoms known as Horner's syndrome:

  • [caption id="attachment_" align="alignright" width=""]Fig - Left sided Horner's syndrome. Note the partial ptosis.Fig - Left sided Horner's syndrome. Note the partial ptosis.[/caption]

    Partial Ptosis - drooping of the upper eyelid. This is due to paralysis of the superior tarsal muscle, which acts to help open the eyelid.

  • Miosis - constriction of the pupil. This is due to paralysis of the dilator pupillae, a muscle located within the eye that acts to dilate the pupil.
  • Anhidrosis - decreased sweating (affecting the same side of the face as the lesion). This is due to a loss of innervation to the sweat glands of the face.

Horner's syndrome has a multitude of causes. These include spinal cord lesions, traumatic injury and a Pancoast tumour (a cancer affecting the apex of the lung, which can involve the ganglia).

[end-clinical]

Summary Table

GangliaVertebral LevelArteries InvolvedEffector Organ(s)
Superior cervical ganglionC1-C4Common, external and internal carotid arteries
  • Eyeball
  • Face
  • Nasal glands
  • Pharynx
  • Glands of the palate and nasal cavity
  • Salivatory glands
  • Lacrimal glands
  • Sweat glands
  • Pineal gland
  • Dilator pupillae
  • Superior tarsal muscle
  • Carotid body
  • Heart
  • Arterial smooth muscle
Middle cervical ganglionC6Inferior thyroid artery
  • Larynx
  • Trachea
  • Pharynx
  • Upper oesophagus
  • Heart
  • Arterial smooth muscle
Inferior cervical ganglionC7Vertebral and subclavian arteries
  • Heart
  • Arterial smooth muscle
Sours: https://teachmeanatomy.info/head/nerves/sympathetic/

Now discussing:

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