Increase, Oxidative Stress

NOCAS

Cigarette smoke

DTXSID5035038

60-35-5

DLFVBJFMPXGRIB-UHFFFAOYSA-N

Acetamide

Acetamid acetamida Acetic acid amide Acetimidic acid Ethanamide Ethanimidic acid Methanecarboxamide NSC 25945

DTXSID7020005

103-90-2

RZVAJINKPMORJF-UHFFFAOYSA-N

Acetaminophen

4-Acetamidophenol APAP Paracetamol 4-hydroxyacetanilide Acetamide, N-(4-hydroxyphenyl)- 4-(Acetylamino)phenol 4-(N-Acetylamino)phenol 4-Acetaminophenol 4'-Hydroxyacetanilide Abensanil Acetagesic Acetalgin ACETAMIDE, N-(4-HYDROXYPHENYL) Acetaminofen Acetanilide, 4'-hydroxy- ACETANILIDE, 4-HYDROXY- Algotropyl Alvedon Anaflon Apamide Banesin Ben-u-ron Bickie-mol Biocetamol Cetadol Citramon P Claratal Clixodyne Dafalgan Daphalgan Dial-a-gesic Disprol Doliprane Dolprone Dymadon Efferalgan Endophy Febrilex Febrilix Febro-Gesic Febrolin Fepanil Finimal Gattaphen T Gelocatil Gutte Enteric Homoolan Jin Gang Lestemp Liquagesic Lonarid Lyteca Syrup Minoset Momentum N-(4-Hydroxyphenyl)acetamide N-Acetyl-4-aminophenol N-Acetyl-4-hydroxyaniline N-Acetyl-p-aminophenol Napafen Naprinol Nobedon NSC 109028 NSC 3991 Ortensan p-(Acetylamino)phenol p-Aceaminophenol Pacemol p-Acetamidophenol p-Acetoaminophen P-ACETYLAMINOPHENOL Paldesic panadeine Panadol Panadol Actifast Panadol Extend Panaleve Panasorb Panodil Paracetamol DC Paracetamole Parageniol Paramol Paraspen Parelan Pasolind N Perfalgan Phenaphen Phendon p-Hydroxyacetanilide Prodafalgan Puerxitong Pyrinazine Resfenol Resprin Rhodapop NCR Salzone Tabalgin Tachipirina Tempanal Tralgon Tylenol TylolHot Valadol Valgesic Vermidon Vick Pyrena

DTXSID2020006

968-81-0

VGZSUPCWNCWDAN-UHFFFAOYSA-N

Acetohexamide

Benzenesulfonamide, 4-acetyl-N-[(cyclohexylamino)carbonyl]- 1-(p-Acetylbenzenesulfonyl)-3-cyclohexylurea 1-[(p-Acetylphenyl)sulfonyl]-3-cyclohexylurea Acetohexamid acetohexamida Dimelin Dimelor Dymelor Gamadiabet Hypoglicil Metaglucina Minoral N-(p-Acetylphenylsulfonyl)-N'-cyclohexylurea Ordimel Tsiklamid Urea, 1-[(p-acetylphenyl)sulfonyl]-3-cyclohexyl-

DTXSID7020007

67-66-3

HEDRZPFGACZZDS-UHFFFAOYSA-N

Chloroform

Trichloromethane Methane, trichloro- CARBON TRICHLORIDE Chloroforme cloroformo Formyl trichloride Methane trichloride Methane,trichloro- NSC 77361 Trichloroform UN 1888

DTXSID1020306

110-00-9

YLQBMQCUIZJEEH-UHFFFAOYSA-N

Furan

Divinylene oxide furanne Furfuran Oxacyclopentadiene Tetrole UN 2389

DTXSID6020646

7429-90-5

XAGFODPZIPBFFR-UHFFFAOYSA-N

AZDRQVAHHNSJOQ-UHFFFAOYSA-N

Aluminum

Aisin Metal Fiber Al 050P-H24 ALC Fine Alcan XI 1391 Almi-Paste SSP 303AR Aloxal 3010 Alpaste 00-0506 Alpaste 0100M Alpaste 0100MA Alpaste 0100M-C Alpaste 0200M Alpaste 0200T Alpaste 0230M Alpaste 0230T Alpaste 0241M Alpaste 0300M Alpaste 0500M Alpaste 0539X Alpaste 0620MS Alpaste 0625TS Alpaste 0638-70C Alpaste 0700M Alpaste 0780M Alpaste 0900M Alpaste 100M Alpaste 100MS Alpaste 100MSR Alpaste 1100M Alpaste 1100MA Alpaste 1100N Alpaste 1100NA Alpaste 1109MA Alpaste 1109MC Alpaste 1200M Alpaste 1200T Alpaste 1260MS Alpaste 1500MA Alpaste 1700NL Alpaste 1810YL Alpaste 1830YL Alpaste 1900M Alpaste 1900XS Alpaste 1950M Alpaste 1950N Alpaste 210N Alpaste 2172EA Alpaste 2173 Alpaste 240T Alpaste 241M Alpaste 417 Alpaste 46-046 Alpaste 4-621 Alpaste 4919 Alpaste 50-63 Alpaste 50-635 Alpaste 51-148B Alpaste 51-231 Alpaste 5205N Alpaste 5207N Alpaste 52-509 Alpaste 52-568 Alpaste 5301N Alpaste 5302N Alpaste 53-119 Alpaste 5422NS Alpaste 54-452 Alpaste 54-497 Alpaste 54-542 Alpaste 55-516 Alpaste 55-519 Alpaste 55-574 Alpaste 5620NS Alpaste 5630NS Alpaste 5640NS Alpaste 56-501 Alpaste 5650NS Alpaste 5653NS Alpaste 5654NS Alpaste 5680N Alpaste 5680NS Alpaste 60-600 Alpaste 60-760 Alpaste 60-768 Alpaste 62-356 Alpaste 6340NS Alpaste 6370NS Alpaste 6390NS Alpaste 640NS Alpaste 65-388 Alpaste 66NLB Alpaste 710N Alpaste 7130N Alpaste 7160N Alpaste 7160NS Alpaste 725N Alpaste 740NS Alpaste 7430NS Alpaste 7580NS Alpaste 7620NS Alpaste 7640NS Alpaste 7670M Alpaste 7670NS Alpaste 7675NS Alpaste 7679NS Alpaste 7680N Alpaste 7680NS Alpaste 76840NS Alpaste 7730N Alpaste 7770N Alpaste 7830N Alpaste 8004 Alpaste 8080N Alpaste 8260NAR Alpaste 891K Alpaste 91-0562 Alpaste 92-0592 Alpaste 93-0595 Alpaste 93-0647 Alpaste 94-2315 Alpaste 95-0570 Alpaste 96-0635 Alpaste 96-2104 Alpaste 97-0510 Alpaste 97-0534 Alpaste AW 520B Alpaste AW 612 Alpaste AW 9800 Alpaste F 795 Alpaste FM 7680K Alpaste FX 440 Alpaste FX 910 Alpaste FZ 0534 Alpaste FZU 40C Alpaste G Alpaste HR 8801 Alpaste HS 2 Alpaste J Alpaste K 9800 Alpaste MC 666 Alpaste MC 707 Alpaste MF 20 Alpaste MG 01 Alpaste MG 1000 Alpaste MG 1300 Alpaste MG 500 Alpaste MG 600 Alpaste MH 6601 Alpaste MH 8801 Alpaste MH 9901 Alpaste MR 7000 Alpaste MR 9000 Alpaste MS 630 Alpaste N 1700NL Alpaste NS 7670 Alpaste O 100N Alpaste O 2130 Alpaste O 300M Alpaste P 0100 Alpaste P 1950 Alpaste S Alpaste SAP 110 Alpaste SAP 414P Alpaste SAP 550N Alpaste SCR 5070 Alpaste TCR 2020 Alpaste TCR 2060 Alpaste TCR 2070 Alpaste TCR 3010 Alpaste TCR 3030 Alpaste TCR 3040 Alpaste TCR 3130 Alpaste TD 200T Alpaste UF 500 Alpaste WB 0230 Alpaste WD 500 Alpaste WJP-U 75C Alpaste WX 0630 Alpaste WX 7830 Alpaste WXA 7640 Alpaste WXM 0630 Alpaste WXM 0650 Alpaste WXM 0660 Alpaste WXM 1415 Alpaste WXM 1440 Alpaste WXM 5422 Alpaste WXM 760b Alpaste WXM 7640 Alpaste WXM 7675 Alpaste WXM-T 60B Alpaste WXM-U 75 Alpaste WXM-U 75C Altop X Aluchrome Ultrafin Super Alumat 1600 Alumet H 30 aluminio Aluminium Aluminium Flake Aluminum 27 Aluminum atom Aluminum element Aluminum Flake PCF 7620 Aluminum granules ALUMINUM METAL/GRANULE ALUMINUM PASTE ALUMINUM PIGMENT ALUMINUM TURNINGS Alumi-paste 640NS Alumipaste 91-0562 Alumipaste 98-1822T Alumipaste AW 620 Alumipaste CR 300 Alumipaste GX 180A Alumipaste GX 201A Alumipaste HR 7000 Alumipaste HR 850 Alumipaste MG 11 Alumipaste MH 8801 Aquamet NPW 2900 Aquapaste 205-5 Aquasilver LPW Astroflake 40 Astroflake Black N 020 Astroflake Black N 070 Astroflake LG 40 Astroflake LG 70 Astroflake Silver N 040 Astroshine NJ 1600 Astroshine T 8990 Atomizalumi VA 200 C.I. PIGMENT METAL 1 Chromal IV Chromal X Decomet 1001/10 Decomet 2018/10 Decomet High Gloss Al 1002/10 Ecka AS 081 Eckart 9155 Eterna Brite 301-1 Eterna Brite 601-1 Eterna Brite 651-1 Eterna Brite EBP 251PA Eterna Brite Primier 251PA Ferro FX 53-038 Friend Color F 500GR-W Friend Color F 500WT Friend Color F 700RE-W Friend Color F 701RE-W Hi Print 60T High Print 60T Hisparkle HS 2 Hydro Paste 8726 Hydrolac WHH 2153 Hydrolan 3560 Hydrolux Reflexal 100 Hydroshine WS 1001 JISA 51010P Kryal Z Lansford 243 LE Sheet 800 Leafing Alpaste LG-H Silver 25 Lunar Al-V 95 Metallux 161 Metallux 2154 Metallux 2192 Metalure Metalure 55350 Metalure L 55350 Metalure L 59510 Metalure W 2001 Metapor Metasheen 1800 Metasheen HR 0800 Metasheen KM 100 Metasheen KM 1000 Metasheen Slurry 1807 Metasheen Slurry 1811 Metasheen Slurry KM 100 Metax G Metax S Mirror Glow 1000 Mirror Glow 600 Mirrorsheen Noral Aluminium Noral Ink Grade Aluminium Obron 10890 Offset FM 4500 Puratronic Reflexal 145 Reynolds 400 Reynolds 4-301 Reynolds 4-591 Reynolds 667 SAP 260PW-HS SAP-FM 4010 SBC 516-20Z Scotchcal 7755SE Serumekku Setanium 50MIS-H8 Siberline ET 2025 Siberline ST 21030E1 Silvar A Silver VT 522 Silverline SSP 353 Silvex 793-20C Sparkle Silver 3141ST Sparkle Silver 3500 Sparkle Silver 3641 Sparkle Silver 5000AR Sparkle Silver 516AR Sparkle Silver 5242AR Sparkle Silver 5245AR Sparkle Silver 5271AR Sparkle Silver 5500 Sparkle Silver 5745 Sparkle Silver 7000AR Sparkle Silver 7005AR Sparkle Silver 7500 Sparkle Silver 960-25E1 Sparkle Silver E 1745AR Sparkle Silver L 1526AR Sparkle Silver Premier 751 Sparkle Silver SS 3130 Sparkle Silver SS 5242AR Sparkle Silver SS 5588 Sparkle Silver SSP 132AR Special PCR 507 Splendal 6001BG Spota Mobil 801 SSP 760-20C Stapa Aloxal PM 2010 Stapa Aloxal PM 3010 Stapa Aloxal PM 4010 Stapa Hydrolac BG 8n.1 Stapa Hydrolac BGH Chromal X Stapa Hydrolac PM Chromal VIII Stapa Hydrolac W 60NL Stapa Hydrolac WH 16 Stapa Hydrolac WH 66NL Stapa Hydrolux 2192 Stapa Hydrolux 8154 Stapa IL Hydrolan 2192-55900G Stapa Metallic R 607 Stapa Metallux 1050 Stapa Metallux 211 Stapa Metallux 212 Stapa Metallux 2196 Stapa Metallux 274 Stapa Mobilux 181 Stapa Offset 3000 Stapa PV 10 Stapa VP 46432G Starbrite 2100 Super Fine 18000 Super Fine 22000 Supramex 2022 Toyo Aluminum 02-0005 Toyo Aluminum 93-3040 Transmet K 102HE Tufflake 3645 Tufflake 5843 UN 1396 US Aluminum 809 Valimet H 2 Valimet H 3 White Silver 7080N White Silver 7130N

DTXSID3040273

7440-43-9

BDOSMKKIYDKNTQ-UHFFFAOYSA-N

Cadmium

Cadimium CADMIUM BLUE CADMIUM, IN PLATTEN, STANGEN, BROCKEN,KOERNER

DTXSID1023940

7439-97-6

QSHDDOUJBYECFT-UHFFFAOYSA-N

Mercury

Liquid silver Mercure MERCURIC METAL TRIPLE DISTILLED mercurio Mercury element Quecksilber Quicksilver UN 2024 UN 2809

DTXSID1024172

7440-61-1

JFALSRSLKYAFGM-UHFFFAOYSA-N

Uranium

Uranium, isotope of mass 238 238U Element UN 2979 (DOT) Uranium I

DTXSID1042522

7440-38-2

RQNWIZPPADIBDY-UHFFFAOYSA-N

Arsenic

As Arsenic black ARSENIC METAL arsenico Grey arsenic UN 1558

DTXSID4023886

7440-22-4

BQCADISMDOOEFD-UHFFFAOYSA-N

Silver

Ag Nanopaste NPS-J 90 Ag Sphere 2 Ag-C-GS Algaedyn Arctic Silver 3 Argentum Astroflake 5 Carey Lea silver Colloidal silver Dotite XA 208 Du Pont 4943 ECM 100AF4810 Enlight 600 Enlight silver plate 600 Epinall Finesphere SVND 102 Fordel DC FP 5369-502 Jelcon SH 1 Jungindai Takasago 300 KS (metal) LCP 1-19SFS Metz 3000-1 Nanomelt AGC-A Nanomelt Ag-XA 301 Nanomelt Ag-XF 301 Nanomelt Ag-XF 301H Nanopaste NPS-J 90 Perfect Silver Puff Silver X 1200 RT 1710S-C1 SD (metal) Shell Silver Silbest E 20 Silbest F 20 Silbest J 18 Silbest TC 12 Silbest TC 20E Silbest TC 25A Silbest TCG 1 Silbest TCG 7 Silcoat AgC 103 Silcoat AgC 2011 Silcoat AgC 209 Silcoat AgC 2190 Silcoat AgC 222 Silcoat AgC 2411 Silcoat AgC 74T Silcoat AgC-A Silcoat AgC-AO Silcoat AgC-B Silcoat AgC-BO Silcoat AgC-D Silcoat AgC-G Silcoat AgC-GS Silcoat AgC-L Silcoat AgC-O Silcoat GS Silcoat RF 200 Silflake 135 Silsphere 514 Silver atom Silver element Silver Flake 1 Silver Flake 25 Silver Flake 52 Silver Flake 7A SILVER FLAKES Silver metal Silvest TCG 11N Technic 299 Technic 450 Techno Alpha 175

DTXSID4024305

7439-96-5

PWHULOQIROXLJO-UHFFFAOYSA-N

Manganese

Colloidal manganese Cutaval Manganese element Manganese fulleride Manganese metal alloy Manganese-55 manganeso

DTXSID2024169

7440-02-0

PXHVJJICTQNCMI-UHFFFAOYSA-N

Nickel

Carbonyl 255 Carbonyl Ni 123 Carbonyl Ni 283 Carbonyl Nickel 123 Carbonyl Nickel 283 Carbonyl Nickel 287 Cerac N 2003 CNS 10 Micron Exmet 4 Ni X-4/0 Fibrex P Incofoam Nickel element NICKEL ROUND ANODES Nicrobraz LM:BNi 2 Ni-Flake 95 Novamet 123 Novamet 4SP Novamet 4SP10 Novamet 525 Novamet CNS 400 Novamet HCA 1 Novamet NI 255 Raney nickel Raney nickel 2800 UN 1325 UN 2881

DTXSID2020925

7440-66-6

HCHKCACWOHOZIP-UHFFFAOYSA-N

Zinc

Zn Asarco L 15 C.I. Pigment Black 16 Merrillite NC-Zinc Rheinzink Stapa TE Zinc AT UF (metal) UN 1436 Zinc dust Zinc Dust 3 Zinc Dust 500 mesh Zinc Dust LS 2 Zinc Dust MCS Zinc Flakes GTT ZINC METAL ZINC MOSSY ZINC STRIP ZINC, MOSSY Zincsalt GTT

DTXSID7035012

75-07-0

IKHGUXGNUITLKF-UHFFFAOYSA-N

Acetaldehyde

Ethanal acetaldehido Acetaldehyd Acetic aldehyde Ethyl aldehyde NSC 7594 UN 1089

DTXSID5039224

107-02-8

HGINCPLSRVDWNT-UHFFFAOYSA-N

Acrolein

2-Propenal 2-Propen-1-al 2-Propen-1-one acrilaldehido Acroleina Acrylaldehyd Acrylaldehyde Acrylic aldehyde Allyl aldehyde Aqualin Magnacide B Magnacide H NSC 8819 Prop-2-en-1-al Propenal UN 1092

DTXSID5020023

54-11-5

SNICXCGAKADSCV-JTQLQIEISA-N

Nicotine

Nicotine alkaloid Pyridine, 3-[(2S)-1-methyl-2-pyrrolidinyl]- (-)-(S)-Nicotine (-)-3-(1-Methyl-2-pyrrolidyl)pyridine (-)-Nicotine (-)-β-Pyridyl-α-N-methylpyrrolidine (S)-(-)-Nicotine (S)-3-(1-Methyl-2-pyrrolidinyl)pyridine (S)-Nicotine 3-[(2S)-1-Methyl-2-pyrrolidinyl]pyridine Flux Maag Habitrol l-Nicotine Nicabate Nicoderm Nicolan Niconil Nicopatch Nicorette Nicotell TTS Nicotin nicotina NICOTINE, Nicotinell Nicotrol NSC 5065 Pyridine, 3-(1-methyl-2-pyrrolidinyl)-, (S)- Tabazur UN 1654 XL All Insecticide

DTXSID1020930

10028-15-6

CBENFWSGALASAD-UHFFFAOYSA-N

Ozone

Atmospheric ozone Healozone Oxygen, mol. Ozone(16O16O16O) Triatomic oxygen

DTXSID0021098

7446-09-5

RAHZWNYVWXNFOC-UHFFFAOYSA-N

Sulfur dioxide

SO2 dioxido de azufre Dioxyde de soufre Fermenticide liquid Schwefeldioxid SULFUR DIOXIDE, LIQUID Sulfur superoxide Sulfurous acid anhydride Sulfurous anhydride Sulfurous oxide sulphur dioxide UN 1079

DTXSID6029672

50-00-0

WSFSSNUMVMOOMR-UHFFFAOYSA-N

Formaldehyde

Fannoform Floguard 1015 formaldehido Formaldehyd Formalin LM Formalin Taisei Formalith Formic aldehyde Lysoform Methaldehyde Methanal Methyl aldehyde Methylene oxide Morbicid NSC 298885 Optilyse Oxomethane Oxymethylene Superlysoform UN 1198 UN 2209 Caswell No. 465 EINECS 200-001-8 EPA Pesticide Chemical Code 043001 Formaldehyde, gas Formalin 40 NCI-C02799 RCRA waste number U122 Aldehyd mravenci Aldehyde formique Aldeide formica Formalin-loesungen Oplossingen UNII-1HG84L3525 Formalaz

DTXSID7020637

10102-43-9

MWUXSHHQAYIFBG-UHFFFAOYSA-N

Nitric oxide

Nitric oxide (NO) Amidogen, oxo- monoxido de nitrogeno Monoxyde d'azote Nitric oxide trimer Nitrogen monooxide nitrogen monoxide Nitrogen(II) oxide Nitrosyl radical Oxido nitrico Stickstoffmonoxid UN 1660

DTXSID1020938

NOCAS

Diesel engine exhaust

Diesel Exhaust DE

DTXSID1024043

PR:000007628

PR forkhead box protein J1

GO:0031514

GO motile cilium

MP:0003674

MP oxidative stress

HP:0012262

HP Abnormal ciliary motility

VT:0001947

VT mucociliary clearance trait

VT:0002327

VT respiratory function trait

WIKI increased

WIKI decreased

WIKI occurrence

Cigarette smoke

2021-06-24T07:10:58

2021-09-28T09:07:54

Acetaminophen

2016-11-29T18:42:26

Chloroform

2016-11-29T18:42:27

furan

2020-05-01T14:35:22

Platinum

2022-02-04T14:36:54

Aluminum

2022-02-04T14:42:11

Cadmium

2017-10-25T08:33:12

Mercury

2016-11-29T18:42:19

Uranium

2021-08-05T14:28:50

Arsenic

2021-04-27T00:15:21

Silver

2022-02-03T11:20:11

Manganese

2022-02-04T14:47:23

Nickel

2022-02-04T14:47:59

Zinc

2022-02-04T15:05:00

nanoparticles

2016-12-21T09:40:06

Irradiation

2021-07-23T07:21:12

Acetaldehyde

2021-07-22T07:51:43

Acrolein

2017-08-15T09:55:59

2021-09-28T08:23:20

Nicotine

2021-07-22T07:53:07

Ozone

2021-07-21T10:18:56

2021-09-28T08:26:52

Sex hormone

2021-09-28T07:57:44

Sulfur dioxide

2021-07-22T09:49:34

Formaldehyde

2021-07-22T09:51:10

PM10

2021-07-22T09:54:46

Nitric oxide

2021-07-22T09:57:38

Diesel engine exhaust

2021-08-06T08:41:15

2021-09-28T08:55:47

WikiUser_26

ApacheUser rodents

9606

NCBI Homo sapiens

WCS_7955

common ecological species Danio rerio

WCS_8355

common ecological species Xenopus laevis

10090

NCBI Mus musculus

10116

NCBI Rattus norvegicus

9986

NCBI Oryctolagus cuniculus

WCS_9913

common ecological species Bos taurus

10141

NCBI Cavia porcellus

8400

NCBI Lithobates catesbeianus

9825

NCBI Sus scrofa domesticus

9940

NCBI Ovis aries

9612

NCBI Canis lupus

WCS_8400

common ecological species Rana catesbeiana

WCS_9606

common toxicological species human

79327

NCBI Schmidtea mediterranea

Increase, Oxidative Stress

Molecular

Oxidative stress is defined as an imbalance in the production of reactive oxygen species (ROS) and antioxidant defenses. High levels of oxidizing free radicals can be very damaging to cells and molecules within the cell.  As a result, the cell has important defense mechanisms to protect itself from ROS. For example, Nrf2 is a transcription factor and master regulator of the oxidative stress response. During periods of oxidative stress, Nrf2-dependent changes in gene expression are important in regaining cellular homeostasis (Nguyen, et al., 2009) and can be used as indicators of the presence of oxidative stress in the cell.  In addition to the directly damaging actions of ROS, cellular oxidative stress also changes cellular activities on a molecular level. Redox sensitive proteins have altered physiology in the presence and absence of ROS, which is caused by the oxidation of sulfhydryls to disulfides on neighboring amino acids (Antelmann & Helmann 2011). Importantly Keap1, the negative regulator of Nrf2, is regulated in this manner (Itoh, et al. 2010).  ROS also undermine the mitochondrial defense system from oxidative damage. The antioxidant systems consist of superoxide dismutase, catalase, glutathione peroxidase and glutathione reductase, as well as antioxidants such as α-tocopherol and ubiquinol, or antioxidant vitamins and minerals including vitamin E, C, carotene, lutein, zeaxanthin, selenium, and zinc (Fletcher, 2010). The enzymes, vitamins and minerals catalyze the conversion of ROS to non-toxic molecules such as water and O2. However, these antioxidant systems are not perfect and endogenous metabolic processes and/or exogenous oxidative influences can trigger cumulative oxidative injuries to the mitochondria, causing a decline in their functionality and efficiency, which further promotes cellular oxidative stress (Balasubramanian, 2000; Ganea & Harding, 2006; Guo et al., 2013; Karimi et al., 2017).   However, an emerging viewpoint suggests that ROS-induced modifications may not be as detrimental as previously thought, but rather contribute to signaling processes (Foyer et al., 2017).    Sources of ROS Production  Direct Sources: Direct sources involve the deposition of energy onto water molecules, breaking them into active radical species. When ionizing radiation hits water, it breaks it into hydrogen (H*) and hydroxyl (OH*) radicals by destroying its bonds. The hydrogen will create hydroxyperoxyl free radicals (HO2*) if oxygen is available, which can then react with another of itself to form hydrogen peroxide (H2O2) and more O2 (Elgazzar and Kazem, 2015). Antioxidant mechanisms are also affected by radiation, with catalase (CAT) and peroxidase (POD) levels rising as a result of exposure (Seen et al. 2018; Ahmad et al. 2021).   Indirect Sources: An indirect source of ROS is the mitochondria, which is one of the primary producers in eukaryotic cells (Powers et al., 2008).  As much as 2% of the electrons that should be going through the electron transport chain in the mitochondria escape, allowing them an opportunity to interact with surrounding structures. Electron-oxygen reactions result in free radical production, including the formation of hydrogen peroxide (H2O2) (Zhao et al., 2019). The electron transport chain, which also creates ROS, is activated by free adenosine diphosphate (ADP), O2, and inorganic phosphate (Pi) (Hargreaves et al. 2020; Raimondi et al. 2020; Vargas-Mendoza et al. 2021). The first and third complexes of the transport chain are the most relevant to mammalian ROS production (Raimondi et al., 2020). The mitochondria has its own set of DNA and it is a prime target of oxidative damage (Guo et al., 2013). ROS is also produced through nicotinamide adenine dinucleotide phosphate oxidase (Nox) stimulation, an event commenced by angiotensin II, a product/effector of the renin-angiotensin system (Nguyen Dinh Cat et al. 2013; Forrester et al. 2018). Other ROS producers include xanthine oxidase, immune cells (macrophage, neutrophils, monocytes, and eosinophils), phospholipase A2 (PLA2), monoamine oxidase (MAO), and carbon-based nanomaterials (Powers et al. 2008; Jacobsen et al. 2008; Vargas-Mendoza et al. 2021).

Oxidative Stress: Direct measurement of ROS is difficult because ROS are unstable. The presence of ROS can be assayed indirectly by measurement of cellular antioxidants, or by ROS-dependent cellular damage. Listed below are common methods for detecting the KE, however there may be other comparable methods that are not listed  <ul> <li>Detection of ROS by chemiluminescence (https://www.sciencedirect.com/science/article/abs/pii/S0165993606001683) </li> <li>Detection of ROS by chemiluminescence is also described in OECD TG 495 to assess phototoxic potential. </li> <li>Glutathione (GSH) depletion. GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green- ab138881.html). </li> <li>TBARS. Oxidative damage to lipids can be measured by assaying for lipid peroxidation using TBARS (thiobarbituric acid reactive substances) using a commercially available kit. </li> <li>8-oxo-dG. Oxidative damage to nucleic acids can be assayed by measuring 8-oxo-dG adducts (for which there are a number of ELISA based commercially available kits),or HPLC, described in Chepelev et al. (Chepelev, et al. 2015). </li> </ul>    Molecular Biology: Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assay for Nrf2 activity include:  <ul> <li>Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus Western blot for increased Nrf2 protein levels </li> <li>Western blot of cytoplasmic and nuclear fractions to observe translocation of Nrf2 protein from the cytoplasm to the nucleus qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences) </li> <li>Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway (e.g., Jackson et al. 2014) </li> <li>OECD TG422D describes an ARE-Nrf2 Luciferase test method </li> </ul> In general, there are a variety of commercially available colorimetric or fluorescent kits for detecting Nrf2 activation. <table border="1"> <tbody> <tr> <td> Assay Type & Measured Content  </td> <td> Description  </td> <td> Dose Range Studied  </td> <td> Assay Characteristics (Length/Ease of use/Accuracy)  </td> </tr> <tr> <td> ROS  Formation in the Mitochondria assay (Shaki et al., 2012)  </td> <td> “The mitochondrial ROS measurement was performed flow cytometry using DCFH-DA. Briefly, isolated kidney mitochondria were incubated with UA (0, 50, 100 and 200 µM) in respiration buffer containing (0.32 mM sucrose, 10mM Tris, 20 mM Mops, 50 µM EGTA, 0.5 mM MgCl2, 0.1 mM KH2PO4 and 5 mM sodium succinate) [32]. In the interval times of 5, 30 and 60 min following the UA addition, a sample was taken and DCFH-DA was added (final concentration, 10 µM) to mitochondria and was then incubated for 10 min.Uranyl acetate-induced ROS generation in isolated kidney mitochondria were determined through the flow cytometry (Partec, Deutschland) equipped with a 488-nm argon ion laser and supplied with the Flomax software and the signals were obtained using a 530-nm bandpass filter (FL-1 channel). Each determination is based on the mean fluorescence intensity of 15,000 counts.”    </td> <td> 0, 50,100 and 200 µM of Uranyl Acetate    </td> <td>  Long/ Easy High accuracy    </td> </tr> <tr> <td> Mitochondrial Antioxidant Content Assay Measuring GSH content (Shaki et al., 2012)    </td> <td> “GSH content was determined using DTNB as the indicator and spectrophotometer method for the isolated mitochondria. The mitochondrial fractions (0.5 mg protein/ml) were incubated with various concentrations of uranyl acetate for 1 h at 30 °C and then 0.1 ml of mitochondrial fractions was added into 0.1 mol/l of phosphate buffers and 0.04% DTNB in a total volume of 3.0 ml (pH 7.4). The developed yellow color was read at 412 nm on a spectrophotometer (UV-1601 PC, Shimadzu, Japan). GSH content was expressed as µg/mg protein.”  </td> <td> 0, 50,  100, or  200 µM  Uranyl Acetate  </td> <td>   </td> </tr> <tr> <td> H2O2 Production Assay Measuring H2O2 Production in isolated mitochondria (Heyno et al., 2008)    </td> <td> “Effect of CdCl2 and antimycin A (AA) on H2O2 production in isolated mitochondria from potato. H2O2 production was measured as scopoletin oxidation. Mitochondria were incubated for 30 min in the measuring buffer  (see the Materials and Methods) containing 0.5 mM succinate as an electron donor and 0.2 µM mesoxalonitrile 3‐chlorophenylhydrazone (CCCP) as an uncoupler, 10 U horseradish peroxidase and 5 µM scopoletin.”   </td> <td> 0, 10, 30  µM Cd2+     2 µM antimycin A  </td> <td>   </td> </tr> <tr> <td> Flow Cytometry ROS & Cell Viability (Kruiderig et al., 1997)    </td> <td> “For determination of ROS, samples taken at the indicated time points were directly transferred to FACScan tubes. Dih123 (10 mM, final concentration) was added and cells were incubated at 37°C in a humidified atmosphere (95% air/5% CO2) for 10 min. At t 5 9, propidium iodide (10 mM, final concentration) was added, and cells were analyzed by flow cytometry at 60 ml/min. Nonfluorescent Dih123 is cleaved by ROS to fluorescent R123 and detected by the FL1 detector as described above for Dc (Van de Water 1995)”“For determination of ROS, samples taken at the indicated time points were directly transferred to FACScan tubes. Dih123 (10 mM, final concentration) was added and cells were incubated at 37°C in a humidified atmosphere (95% air/5% CO2) for 10 min. At t 5 9, propidium iodide (10 mM, final concentration) was added, and cells were analyzed by flow cytometry at 60 ml/min. Nonfluorescent Dih123 is cleaved by ROS to fluorescent R123 and detected by the FL1 detector as described above for Dc (Van de Water 1995)”  </td> <td>   </td> <td>           Strong/easy medium  </td> </tr> <tr> <td> DCFH-DA  Assay Detection of hydrogen peroxide production (Yuan et al.,  2016)  </td> <td> Intracellular ROS production was measured using DCFH-DA as a probe. Hydrogen peroxide oxidizes DCFH to DCF. The probe is hydrolyzed intracellularly to DCFH carboxylate anion. No direct reaction with H2O2 to form fluorescent production.    </td> <td> 0-400  µM  </td> <td> Long/ Easy High accuracy  </td> </tr> <tr> <td> H2-DCF-DAAssay Detection of superoxide production (Thiebault etal., 2007)    </td> <td> This dye is a stable nonpolar compound which diffuses readily into the cells and yields H2-DCF. Intracellular OH or ONOO- react with H2-DCF when cells contain peroxides, to form the highly fluorescent compound DCF, which effluxes the cell. Fluorescence intensity of DCF is measured using a fluorescence spectrophotometer.  </td> <td> 0–600  µM  </td> <td> Long/ Easy High accuracy  </td> </tr> <tr> <td> CM-H2DCFDA  Assay (Eruslanov  & Kusmartsev, 2009)  </td> <td> The dye (CM-H2DCFDA) diffuses into the cell and is cleaved by esterases, the thiol reactive chlormethyl group reacts with intracellular glutathione which can be detected using flow cytometry.  </td> <td>   </td> <td> Long/Easy/ High Accuracy  </td> </tr> </tbody> </table>   <table border="1"> <tbody> <tr> <td> Method of Measurement   </td> <td> References   </td> <td> Description   </td> <td colspan="2"> OECD-Approved Assay  </td> </tr> <tr> <td> Chemiluminescence   </td> <td> (Lu, C. et al., 2006;   Griendling, K. K., et al., 2016)  </td> <td> ROS can induce electron transitions in molecules, leading to electronically excited products. When the electrons transition back to ground state, chemiluminescence is emitted and can be measured. Reagents such as luminol and lucigenin are commonly used to amplify the signal.   </td> <td colspan="2"> No    </td> </tr> <tr> <td> Spectrophotometry   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> NO has a short half-life. However, if it has been reduced to nitrite (NO2-), stable azocompounds can be formed via the Griess Reaction, and further measured by spectrophotometry.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Direct or Spin Trapping-Based electron paramagnetic resonance (EPR) Spectroscopy   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> The unpaired electrons (free radicals) found in ROS can be detected with EPR and is known as electron paramagnetic resonance. A variety of spin traps can be used.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Nitroblue Tetrazolium Assay   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> The Nitroblue Tetrazolium assay is used to measure O2.− levels. O2.− reduces nitroblue tetrazolium (a yellow dye) to formazan (a blue dye), and can be measured at 620 nm.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Fluorescence analysis of dihydroethidium (DHE) or Hydrocyans   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> Fluorescence analysis of DHE is used to measure O2.− levels.  O2.− is reduced to O2 as DHE is oxidized to 2-hydroxyethidium, and this reaction can be measured by fluorescence. Similarly, hydrocyans can be oxidized by any ROS, and measured via fluorescence.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Amplex Red Assay   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> Fluorescence analysis to measure extramitochondrial or extracellular H2O2 levels. In the presence of horseradish peroxidase and H2O2, Amplex Red is oxidized to resorufin, a fluorescent molecule measurable by plate reader.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Dichlorodihydrofluorescein Diacetate (DCFH-DA)   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> An indirect fluorescence analysis to measure intracellular H2O2 levels.  H2O2 interacts with peroxidase or heme proteins, which further react with DCFH, oxidizing it to dichlorofluorescein (DCF), a fluorescent product.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> HyPer Probe   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> Fluorescent measurement of intracellular H2O2 levels. HyPer is a genetically encoded fluorescent sensor that can be used for in vivo and in situ imaging.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Cytochrome c Reduction Assay   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> The cytochrome c reduction assay is used to measure O2.− levels. O O2.− is reduced to O2 as ferricytochrome c is oxidized to ferrocytochrome c, and this reaction can be measured by an absorbance increase at 550 nm.   </td> <td colspan="2"> No  </td> </tr> <tr> <td> Proton-electron double-resonance imaging (PEDRI)   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> The redox state of tissue is detected through nuclear magnetic resonance/magnetic resonance imaging, with the use of a nitroxide spin probe or biradical molecule.   </td> <td colspan="2"> No              </td> </tr> <tr> <td> Glutathione (GSH) depletion   </td> <td> (Biesemann, N. et al., 2018)   </td> <td> A downstream target of the Nrf2 pathway is involved in GSH synthesis. As an indication of oxidation status, GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., <a href="http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html" rel="noreferrer noopener" target="_blank">http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html</a>).    </td> <td colspan="2"> No  </td> </tr> <tr> <td> Thiobarbituric acid reactive substances (TBARS)   </td> <td> (Griendling, K. K., et al., 2016)  </td> <td> Oxidative damage to lipids can be measured by assaying for lipid peroxidation with TBARS using a commercially available kit.    </td> <td colspan="2"> No  </td> </tr> <tr> <td> Protein oxidation (carbonylation)  </td> <td> (Azimzadeh et al., 2017; Azimzadeh et al., 2015; Ping et al., 2020)  </td> <td> Can be determined with ELISA or a commercial assay kit. Protein oxidation can indicate the level of oxidative stress.  </td> <td colspan="2"> No  </td> </tr> <tr> <td> Seahorse XFp Analyzer  </td> <td> Leung et al. 2018  </td> <td> The Seahorse XFp Analyzer provides information on mitochondrial function, oxidative stress, and metabolic dysfunction of viable cells by measuring respiration (oxygen consumption rate; OCR) and extracellular pH (extracellular acidification rate; ECAR).  </td> <td> No  </td> </tr> </tbody> </table>   Molecular Biology: Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assays for Nrf2 activity include:   <table border="1"> <tbody> <tr> <td> Method of Measurement   </td> <td> References   </td> <td> Description   </td> <td> OECD-Approved Assay  </td> </tr> <tr> <td> Immunohistochemistry   </td> <td> (Amsen, D., de Visser, K. E., and Town, T., 2009)  </td> <td> Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus    </td> <td> No  </td> </tr> <tr> <td> qPCR   </td> <td> (Forlenza et al., 2012)  </td> <td> qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences)   </td> <td> No  </td> </tr> <tr> <td> Whole transcriptome profiling via microarray or via RNA-seq followed by a pathway analysis  </td> <td> (Jackson, A. F. et al., 2014)  </td> <td> Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway  </td> <td> No  </td> </tr> </tbody> </table>

Taxonomic applicability: Occurrence of oxidative stress is not species specific.   Life stage applicability: Occurrence of oxidative stress is not life stage specific.  Sex applicability: Occurrence of oxidative stress is not sex specific.  Evidence for perturbation by prototypic stressor: There is evidence of the increase of oxidative stress following perturbation from a variety of stressors including exposure to ionizing radiation and altered gravity (Bai et al., 2020; Ungvari et al., 2013; Zhang et al., 2009).

High Mixed

High

All life stages

High High

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2026-02-11T07:05:27

FOXJ1 Protein, Decreased

Cellular

The epithelium of the respiratory tract has a powerful defense mechanism against air-borne pollutants due to the combined performance of mucus-producing goblet cells and ciliated cells that are covered with microtubule-based projections, the cilia. In response to various irritants and pathogens mucus is secreted by goblet cells, and cilia sweep mucus upward by coordinated beating motions thus clearing the airways from these substances. The ciliated airway epithelial cells are typically covered by hundreds of motile cilia. Cilia formation is initiated and coordinated by a distinct gene expression program, led by the transcription factor forkhead box J1 (FOXJ1) (Brody et al., 2000; Zhou and Roy, 2015). In addition to the respiratory tract, FOXJ1 is expressed also in the ciliated cells of the reproductive and central nervous systems (Blatt et al., 1999; Hackett et al., 1995; Lim et al., 1997).  The multiple motile cilia assembly factors MCIDAS and GMNC converge in positive regulation of FOXJ1 (Arbi et al., 2016; Berta et al., 2016; Stubbs et al., 2012), whereas NOTCH signaling, IL-13-or EGF (epidermal growth factor)-triggered signaling antagonize FOXJ1-driven multiciliogenesis (Gerovac and Fregien, 2016; Gerovac et al., 2014; Gomperts et al., 2007; Shaykhiev et al., 2013). Various other factors are involved in multiple motile cilia assembly, including MYB (acts early in multiciliogenesis downstream of MCIDAS), RFX3 (can act as a co-factor for FOXJ1), ULK4 (modulates the expression of FOXJ1), Wnt signaling, etc. (Choksi et al., 2014; Liu et al., 2016; Schmid et al., 2017; Tan et al., 2013). Most of these factors act upstream or parallel to FOXJ1. FOXJ1 appears to be the major factor in multiciliogenesis, whereby its activity is necessary and also sufficient for programming cells to assemble functional motile cilia (Vij et al., 2012). FOXJ1 is a master regulator of motile ciliogenesis and is essential to program cells to grow motile cilia (Zhou and Roy, 2015). This key event represents the decrease in the levels or absence of FOXJ1 protein in cells of the respiratory tract. The decrease in FOXJ1 levels inhibits ciliogenesis in multiciliated cells of zebrafish and Xenopus (Stubbs et al., 2008). The knockdown of FOXJ1 results in almost complete absence of cilia in mouse epithelial cells (Brody et al., 2000; Chen J. et al., 1998). On the other hand, the overexpression of FOXJ1 rescues cigarette smoke-mediated suppression of cilia growth in human airway epithelium (Brekman et al., 2014).

FOXJ1 protein levels can be measured by Western blot analysis (Brekman et al., 2014; Didon et al., 2013a; Gomperts et al., 2007; Jacquet et al., 2009; Milara et al., 2012), immunofluorescence (Arbi et al., 2016; Gomperts et al., 2007; Valencia-Gattas et al., 2016) or immunohistochemistry (Abedalthagafi et al., 2016; Danielian et al., 2007; Gao et al., 2015). FOXJ1 protein amounts can be inferred from FOXJ1 mRNA levels that can be measured by real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR) (Arbi et al., 2016; Brekman et al., 2014; Didon et al., 2013a; Jacquet et al., 2009; Milara et al., 2012; Stubbs et al., 2012), in situ hybridization (Hackett et al., 1995; Stubbs et al., 2012), and Northern blot analysis (Hackett et al., 1995). In addition, FOXJ1 protein activity can be inferred from FOXJ1 target gene expression levels or from reporter gene expression levels (e.g. luciferase assay) of genes harboring FOXJ1 transcription factor binding sites (Brekman et al., 2014; Lim et al., 1997).

FOXJ1 is functionally conserved throughout diverse groups of metazoans including flatworm Schmidtea mediterranea, zebrafish Danio rerio, African clawed frog Xenopus laevis (Stubbs et al., 2008; Vij et al., 2012; Yu et al., 2008). Ectopic expression of FOXJ1 triggers ciliogenesis in zebrafish and frog (Stubbs et al., 2008; Yu et al., 2008). Overexpression of FOXJ1 transcription factor in the neural tube of a chick induces cilia formation (Cruz C. et al., 2010). There are multiple studies of FOXJ1 in mice and in human cells (Boon et al., 2014; Brekman et al., 2014; Brody et al., 2000; Chen et al., 1998; Choksi et al., 2014). Furthermore, the target genes of FOXJ1, for example RFX3, are regulated by FOXJ1 across different species (Alten et al., 2012; Didon et al., 2013a). FOXJ1 function is important for all life stages from embryo through adulthood (Choksi et al., 2014; Stauber et al., 2017).  FOXJ1 is expressed in the airways of both males and females. In addition to respiratory tract and brain, FOXJ1 is functionally important also in male and female reproductive tissues (Hackett et al., 1995).

UBERON:0000115

UBERON lung epithelium

CL:0005012

CL multi-ciliated epithelial cell

High Mixed

High

All life stages

High High High High

Alten, L., Schuster-Gossler, K., Beckers, A., Groos, S., Ulmer, B., Hegermann, J., et al. (2012). Differential regulation of node formation, nodal ciliogenesis and cilia positioning by Noto and Foxj1. Development 139, 1276-1284. Arbi, M., Pefani, D.E., Kyrousi, C., Lalioti, M.E., Kalogeropoulou, A., Papanastasiou, A.D., et al. (2016). GemC1 controls multiciliogenesis in the airway epithelium. EMBO Rep. 17, 400-413. Azzam, E.I., Jay-Gerin, J.P. and Pain, D. (2012). Ionizing radiation-induced metabolic oxidative stress and prolonged cell injury. Cancer Lett. 327, 48-60. Bernard, M.E., Kim, H., Rajagopalan, M.S., Stone, B., Salimi, U., Rwigema, J.C., et al. (2012). Repopulation of the irradiation damaged lung with bone marrow-derived cells. In Vivo. 26, 9-18. Berta, T., Gabriele, P., Sandra, S.-B., Gabriel, G.-G., A, Y.S., Stephan-Otto, A.C., et al. (2016). GEMC1 is a critical regulator of multiciliated cell differentiation. EMBO J. 35, 942-960. Blatt, E.N., Yan, X.H., Wuerffel, M.K., Hamilos, D.L. and Brody, S.L. (1999). Forkhead transcription factor HFH-4 expression is temporally related to ciliogenesis. Am. J. Respir. Cell Mol. Biol. 21, 168-176. Boon, M., Wallmeier, J., Ma, L., Loges, N.T., Jaspers, M., Olbrich, H., et al. (2014). MCIDAS mutations result in a mucociliary clearance disorder with reduced generation of multiple motile cilia. Nat. Commun. 5, 4418. Brekman, A., Walters, M.S., Tilley, A.E. and Crystal, R.G. (2014). FOXJ1 prevents cilia growth inhibition by cigarette smoke in human airway epithelium in vitro. Am. J. Respir. Cell Mol. Biol. 51, 688-700. Brody, S.L., Yan, X.H., Wuerffel, M.K., Song, S.K. and Shapiro, S.D. (2000). Ciliogenesis and left-right axis defects in forkhead factor HFH-4-null mice. Am. J. Respir. Cell Mol. Biol. 23, 45-51. Chen, J., Knowles, H.J., Hebert, J.L. and Hackett, B.P. (1998). Mutation of the mouse hepatocyte nuclear factor/forkhead homologue 4 gene results in an absence of cilia and random left-right asymmetry. J. Clin. Invest. 102, 1077-1082. Choksi, S.P., Lauter, G., Swoboda, P. and Roy, S. (2014). Switching on cilia: transcriptional networks regulating ciliogenesis. Development 141, 1427-1441. Cruz, C., Ribes, V., Kutejova, E., Cayuso, J., Lawson, V., Norris, D., et al. (2010). Foxj1 regulates floor plate cilia architecture and modifies the response of cells to sonic hedgehog signalling. Development 137, 4271-4282. Didon, L., Zwick, R.K., Chao, I.W., Walters, M.S., Wang, R., Hackett, N.R., et al. (2013). RFX3 Modulation of FOXJ1 regulation of cilia genes in the human airway epithelium. Respir. Res. 14, 70-70. Gerovac, B.J. and Fregien, N.L. (2016). IL-13 inhibits multicilin expression and ciliogenesis via janus kinase/signal transducer and activator of transcription independently of Notch cleavage. Am. J. Respir. Cell Mol. Biol. 54, 554-561. Gerovac, B.J., Valencia, M., Baumlin, N., Salathe, M., Conner, G.E. and Fregien, N.L. (2014). Submersion and hypoxia inhibit ciliated cell differentiation in a notch-dependent manner. Am. J. Respir. Cell Mol. Biol. 51(4), 516-525. Gomperts, B.N., Gong-Cooper, X. and Hackett, B.P. (2004). Foxj1 regulates basal body anchoring to the cytoskeleton of ciliated pulmonary epithelial cells. J. Cell Sci. 117, 1329-1337. Gomperts, B.N., Kim, L.J., Flaherty, S.A. and Hackett, B.P. (2007). IL-13 Regulates Cilia Loss and foxj1 Expression in Human Airway Epithelium. Am. J. Respir. Cell Mol. Biol. 37, 339-346. Hackett, B.P., Brody, S.L., Liang, M., Zeitz, I.D., Bruns, L.A. and Gitlin, J.D. (1995). Primary structure of hepatocyte nuclear factor/forkhead homologue 4 and characterization of gene expression in the developing respiratory and reproductive epithelium. Proc. Natl. Acad. Sci. U. S. A. 92, 4249-4253. Ishikawa, S. and Ito, S. (2017). Repeated whole cigarette smoke exposure alters cell differentiation and augments secretion of inflammatory mediators in air-liquid interface three-dimensional co-culture model of human bronchial tissue. Toxicol. in Vitro 38, 170-178. Koc, M., Taysi, S., Buyukokuroglu, M.E. and Bakan, N. (2003). Melatonin protects rat liver against irradiation-induced oxidative injury. J. Radiat. Res. 44, 211-215. Lim, L., Zhou, H. and Costa, R.H. (1997). The winged helix transcription factor HFH-4 is expressed during choroid plexus epithelial development in the mouse embryo. Proc. Natl. Acad. Sci. U. S. A. 94, 3094-3099. Liu, M., Guan, Z., Shen, Q., Lalor, P., Fitzgerald, U., O'brien, T., et al., 2016. Ulk4 Is essential for ciliogenesis and CSF flow. J. Neurosci. 36, 7589-7600. Milara, J., Armengot, M., Bañuls, P., Tenor, H., Beume, R., Artigues, E., et al. (2012). Roflumilast N-oxide, a PDE4 inhibitor, improves cilia motility and ciliated human bronchial epithelial cells compromised by cigarette smoke in vitro. Brit. J. Pharmacol. 166, 2243-2262. Polosa, R., Emma, R., Cibella, F., Caruso, M., Conte, G., Benfatto, F., et al. (2021). Impact of exclusive e-cigarettes and heated tobacco products use on muco-ciliary clearance. Ther. Adv. Chronic Dis. 12, 20406223211035267-20406223211035267.  Rodrigues-Moreira, S., Moreno, S.G., Ghinatti, G., Lewandowski, D., Hoffschir, F., Ferri, F., et al. (2017). Low-Dose Irradiation Promotes Persistent Oxidative Stress and Decreases Self-Renewal in Hematopoietic Stem Cells. Cell Rep. 20, 3199-3211. Schmid, A., Sailland, J., Novak, L., Baumlin, N., Fregien, N. and Salathe, M. (2017). Modulation of Wnt signaling is essential for the differentiation of ciliated epithelial cells in human airways. FEBS Lett. 591, 3493-3506. Shaykhiev, R., Zuo, W.L., Chao, I., Fukui, T., Witover, B., Brekman, A., et al. (2013). EGF shifts human airway basal cell fate toward a smoking-associated airway epithelial phenotype. Proc. Natl. Acad. Sci. U. S. A. 110, 12102-12107. Shirazi, A., Mihandoost, E., Ghobadi, G., Mohseni, M. and Ghazi-Khansari, M. (2013). Evaluation of radio-protective effect of melatonin on whole body irradiation induced liver tissue damage. Cell J. 14, 292-297. Stauber, M., Weidemann, M., Dittrich-Breiholz, O., Lobschat, K., Alten, L., Mai, M., et al. (2017). Identification of FOXJ1 effectors during ciliogenesis in the foetal respiratory epithelium and embryonic left-right organiser of the mouse. Dev. Biol. 423, 170-188. Stubbs, J.L., Vladar, E.K., Axelrod, J.D. and Kintner, C. (2012). Multicilin promotes centriole assembly and ciliogenesis during multiciliate cell differentiation. Nat. Cell Biol. 14, 140-147. Tan, F.E., Vladar, E.K., Ma, L., Fuentealba, L.C., Hoh, R., Espinoza, F.H., et al. (2013). Myb promotes centriole amplification and later steps of the multiciliogenesis program. Development 140, 4277-4286. Valencia-Gattas, M., Conner, G.E. and Fregien, N.L. (2016). Gefitinib, an EGFR Tyrosine Kinase inhibitor, Prevents Smoke-Mediated Ciliated Airway Epithelial Cell Loss and Promotes Their Recovery. PloS ONE 11, e0160216. Vij, S., Rink, J.C., Ho, H.K., Babu, D., Eitel, M., Narasimhan, V., et al. (2012). Evolutionarily ancient association of the FoxJ1 transcription factor with the motile ciliogenic program. PLoS Genet. 8, e1003019. Yu, X., Ng, C.P., Habacher, H. and Roy, S. (2008). Foxj1 transcription factors are master regulators of the motile ciliogenic program. Nat. Genet. 40, 1445-1453. Zhou, F. and Roy, S. (2015). SnapShot: Motile Cilia. Cell 162, 224-224 e221. AOPWiki 2021-07-23T05:00:42

2021-09-10T04:56:52

Motile Cilia Number/Length, Decreased

Cellular

Motile cilia are microtubule-based organelles that protrude from the cell surface and generate directional flow of fluid with coordinated beating. 50% to 80% of human respiratory epithelium is comprised of ciliated cells covered with multiple motile cilia that move mucus (together with mucus-trapped substances) upward for clearing the airways (Yaghi and Dolovich, 2016). The ciliated airway epithelial cells are typically covered by more than hundred motile cilia (Bustamante-Marin and Ostrowski, 2017). On average, 150 motile cilia were counted per ciliated human epithelial cell in the study by Mao et al. (Mao et al., 2018). In an earlier report, 200 motile cilia per ciliated cell in human trachea is mentioned (Wanner et al., 1996), and, in a more recent study, a range of 100 to 600 ciliary precursors were counted in fully differentiated mouse tracheal epithelial cells correlated with increasing surface area (Nanjundappa et al., 2019). Cilia are 6–7 µm long and 0.2–0.3 µm in diameter (Brooks and Wallingford, 2014; Yaghi and Dolovich, 2016). Ciliated cell density and the motile cilia length and number per cell correlate with ciliary beating frequency which is routinely used as a predictor of the mucociliary clearance efficiency (King, 2006). Morphological changes of airway cilia are expected to impact multiple motile cilia functional integrity. This key event represents the decrease in the numbers or absence of motile cilia or reduction in length of motile cilia.

Acetylated tubulin is a common ciliary marker (Kim et al., 2013; Piperno and Fuller, 1985), and apical acetylated tubulin staining with subsequent microscope image scoring is a frequently used method of cilia detection and enumeration (Johnson et al., 2018; Mao et al., 2018; Stubbs et al., 2008). Staining of beta-tubulin IV, a protein enriched in motile cilia, is another common method of cilia detection (Brekman et al., 2014; Milara et al., 2012). Ciliated cells can also be identified by the presence of axonemal structures on the cell surface using scanning electron microscopy (Gomperts et al., 2007).  Mature cilia numbers could be deduced from ciliary precursors in immunofluorescence assays: ciliary precursors can be calculated from three-dimensional superresolution structured illumination microscopy (3D-SIM) images using e.g. a spot detection tool (Nikon Elements AR 4 Software) (Nanjundappa et al., 2019). For cilia length measurement, the ciliated cells/tissue needs to be stained (Diff-Quik: Dade Behring stain, hematoxylin and eosin staining, labelling with antibodies for ciliary markers such as alpha-tubulin), visualized by microscopy and cilia length quantified (using e.g. ImageJ software or MetaMorph Microscopy Automation & Image Analysis Software) (Brekman et al., 2014; Leopold et al., 2009b; Li et al., 2014). Generally, multiple measurements of one sample and multiple sample preparations of cells/tissues are imaged for reliable quantitation.

The ultrastructural features of human and other mammalian respiratory epithelial cilia and those from lower animals (e.g. flatworms and mollusks) are remarkably similar (Meunier and Azimzadeh, 2016; Wanner et al., 1996). The master regulators of multiciliated cell differentiation, such as NOTCH, GEMC1, MCIDAS, FOXJ1, RFX2/3 are conserved throughout vertebrates (e.g. mammals, Xenopus, zebrafish) and multiple motile cilia across these organisms are functionally similar in generating fluid flow through coordinated beating (Choksi et al., 2014; Meunier and Azimzadeh, 2016; Wessely and Obara, 2008). The motile cilia numbers reach adult levels in the mouse airway epithelium at day 21 after birth (Rawlins et al., 2007; Toskala et al., 2005). At birth, there is no discernable cilia-generated airway fluid flow in mice (Francis et al., 2009). Between postnatal days 3 and 7 the flow is established in trachea correlating with the increase in the density of ciliated cells in the tracheal epithelia (Francis et al., 2009). After airway fluid flow establishment, the KE is applicable to all life stages.

UBERON:0000115

UBERON lung epithelium

CL:0005012

CL multi-ciliated epithelial cell

High Mixed

High

During development and at adulthood

Not Specified High Not Specified Not Specified

Brekman, A., Walters, M.S., Tilley, A.E. and Crystal, R.G. (2014). FOXJ1 prevents cilia growth inhibition by cigarette smoke in human airway epithelium in vitro. Am. J. Respir. Cell Mol. Biol. 51, 688-700. Brooks, E.R. and Wallingford, J.B. (2014). Multiciliated cells. Curr. Biol. 24, R973-982. Bustamante-Marin, X.M. and Ostrowski, L.E. (2017). Cilia and Mucociliary Clearance. Cold Spring Harb. Persp. Biol. 9, a028241. Choksi, S.P., Lauter, G., Swoboda, P. and Roy, S. (2014). Switching on cilia: transcriptional networks regulating ciliogenesis. Development 141, 1427-1441. Francis, R.J., Chatterjee, B., Loges, N.T., Zentgraf, H., Omran, H. and Lo, C.W. (2009). Initiation and maturation of cilia-generated flow in newborn and postnatal mouse airway. Am. J. Physiol. Lung Cell. Mol. Physiol. 296, L1067-1075. Gomperts, B.N., Kim, L.J., Flaherty, S.A. and Hackett, B.P. (2007). IL-13 Regulates Cilia Loss and foxj1 Expression in Human Airway Epithelium. Am. J. Respir. Cell Mol. Biol. 37, 339-346. Hessel, J., Heldrich, J., Fuller, J., Staudt, M.R., Radisch, S., Hollmann, C., et al. (2014). Intraflagellar Transport Gene Expression Associated with Short Cilia in Smoking and COPD. PLoS ONE 9, e85453.  Johnson, J.A., Watson, J.K., Nikolic, M.Z. and Rawlins, E.L. (2018). Fank1 and Jazf1 promote multiciliated cell differentiation in the mouse airway epithelium. Biol. Open 7, bio033944. Kim, G.W., Li, L., Ghorbani, M., You, L. and Yang, X.J. (2013). Mice lacking alpha-tubulin acetyltransferase 1 are viable but display alpha-tubulin acetylation deficiency and dentate gyrus distortion. J. Biol. Chem. 288, 20334-20350. King, M. (2006). Physiology of mucus clearance. Paediatr. Respir Rev. 7, S212-214. Lam, H.C., Cloonan, S.M., Bhashyam, A.R., Haspel, J.A., Singh, A., Sathirapongsasuti, J.F., et al. (2013). Histone deacetylase 6–mediated selective autophagy regulates COPD-associated cilia dysfunction. J. Clin. Invest. 123(12), 5212-5230.  Leopold, P.L., O'mahony, M.J., Lian, X.J., Tilley, A.E., Harvey, B.-G. and Crystal, R.G. (2009). Smoking is associated with shortened airway cilia. PloS ONE 4, e8157. Li, Y.Y., Li, C.W., Chao, S.S., Yu, F.G., Yu, X.M., Liu, J., et al. (2014). Impairment of cilia architecture and ciliogenesis in hyperplastic nasal epithelium from nasal polyps. J. Allergy Clin. Immunol. 134, 1282-1292. Mao, S., Shah, A.S., Moninger, T.O., Ostedgaard, L.S., Lu, L., Tang, X.X., et al. (2018). Motile cilia of human airway epithelia contain hedgehog signaling components that mediate noncanonical hedgehog signaling. Proc. Natl. Acad. Sci. U. S. A. 115, 1370-1375. Meunier, A. and Azimzadeh, J. (2016). Multiciliated Cells in Animals. Cold Spring Harb. Perspect. Biol. 8, a028233. Milara, J., Armengot, M., Bañuls, P., Tenor, H., Beume, R., Artigues, E., et al. (2012). Roflumilast N-oxide, a PDE4 inhibitor, improves cilia motility and ciliated human bronchial epithelial cells compromised by cigarette smoke in vitro. Brit. J. Pharmacol. 166, 2243-2262. Nanjundappa, R., Kong, D., Shim, K., Stearns, T., Brody, S.L., Loncarek, J., et al. (2019). Regulation of cilia abundance in multiciliated cells. Elife 8, e44039.  Piperno, G. and Fuller, M.T. (1985). Monoclonal antibodies specific for an acetylated form of alpha-tubulin recognize the antigen in cilia and flagella from a variety of organisms. J. Cell Biol. 101, 2085-2094. Rawlins, E.L., Ostrowski, L.E., Randell, S.H. and Hogan, B.L. (2007). Lung development and repair: contribution of the ciliated lineage. Proc. Natl. Acad. Sci. U. S. A. 104, 410-417. Simet, S.M., Sisson, J.H., Pavlik, J.A., Devasure, J.M., Boyer, C., Liu, X., et al. (2010). Long-term cigarette smoke exposure in a mouse model of ciliated epithelial cell function. Am. J. Respir. Cell Mol. Biol. 43, 635-640. Stubbs, J.L., Oishi, I., Izpisua Belmonte, J.C. and Kintner, C. (2008). The forkhead protein Foxj1 specifies node-like cilia in Xenopus and zebrafish embryos. Nat. Genet. 40, 1454-1460. Tamashiro, E., Xiong, G., Anselmo-Lima, W.T., Kreindler, J.L., Palmer, J.N., and Cohen, N.A. (2009). Cigarette smoke exposure impairs respiratory epithelial ciliogenesis. Am. J. Rhinol. Allergy 23, 117-122.  Toskala, E., Smiley-Jewell, S.M., Wong, V.J., King, D. and Plopper, C.G. (2005). Temporal and spatial distribution of ciliogenesis in the tracheobronchial airways of mice. Am. J. Physiol. Lung Cell. Mol. Physiol. 289, L454-459. Verra, F., Escudier, E., Lebargy, F., Bernaudin, J.F., De Cremoux, H., and Bignon, J. (1995). Ciliary abnormalities in bronchial epithelium of smokers, ex-smokers, and nonsmokers. Am. J. Respir. Crit. Care Med. 151, 630-634.  Wanner, A., Salathe, M. and O'riordan, T.G. (1996). Mucociliary clearance in the airways. Am. J. Respir. Crit. Care Med. 154, 1868-1902. Wessely, O. and Obara, T. (2008). Fish and frogs: models for vertebrate cilia signaling. Front. Biosci. 13, 1866-1880. Yaghi, A. and Dolovich, M.B. (2016). Airway Epithelial Cell Cilia and Obstructive Lung Disease. Cells. 5, 40. AOPWiki 2021-07-23T06:39:38

2021-09-10T03:24:16

Cilia Beat Frequency, Decreased

CBF, Decreased

Cellular

Cohesive beating of cilia lining the upper and lower respiratory tract is critical for efficient MCC. CBF is influenced by several factors including changes in the physical and chemical properties of the ASL (especially the periciliary fluid), structural modulation in the cilia, concentration of cyclic nucleotides cAMP and cGMP, and intracellular calcium (Ca2+). Formation of cyclic nucleotides such as cGMP is mediated by nitric oxide (NO), which is released by an enzyme family of nitric oxide synthases (NOSs) when the substrate L-arginine (L-Arg) is transformed to L-citrulline. NO activates its receptor protein, soluble guanylate cyclase (sGC), which catalyzes formation of cGMP from guanosine triphosphate (GTP). cGMP then activates protein kinase G (PKG) which has been implicated in the regulation of CBF (Jiao et al., 2011; Li et al., 2000). NO-dependent stimulation of CBF has also been associated with an increase in cAMP-dependent protein kinase A (PKA) (Di Benedetto et al., 1991; Lansley et al., 1992; Salathe et al., 1993; Sanderson and Dirksen, 1989; Schmid et al., 2007; Sisson et al., 1999; Uzlaner and Priel, 1999). An increase in intracellular endogenous cAMP was observed after treatment with isobutyl-1-methylxanthine that also increased CBF (Tamaoki et al., 1989). cAMP accumulation in the airway cilia has been shown to be dependent on Ca2+–calmodulin-dependent PDE1A and indirectly regulates CBF (Kogiso et al., 2018). Increase in CBF after treatment with NO substrate, L-arginine and inhibition of CBF by a NOS inhibitor, N-omega-nitro-L-arginine methyl ester (L-NAME) further provides evidence for the role of NO in increasing CBF (Jiao J. et al., 2011; Sisson J. H., 1995; Uzlaner and Priel, 1999; Yang et al., 1997).  Modulation of CBF is not always accompanied by changes in cAMP levels. PKC activators, phorbol 12-myristate 13-acetate and L-o~-dioctanoylglycerol have been shown to decrease CBF in a concentration- and time-dependent manner in rabbit tracheal epithelial cells (Kobayashi et al., 1989). CBF has been shown to decrease after exposure to inhaled oxidants such as cigarette smoke across different species. A study with 120 subjects showed a significant decrease in nasal CBF following exposure to tobacco smoke (Agius et al., 1998). Exposure to cigarette smoke extract lead to reduction in forskolin-induced CBF in human sinonasal epithelium (Cohen et al., 2009) and  isoproterenol- and methacholine-induced CBF in human adenoid tissues (Wang et al., 2012). This decrease in CBF and unresponsiveness to beta-agonist stimulation occurs in parallel to PKC activation and has been shown to be dependent on the duration of exposure to cigarette smoke in mice (Simet et al., 2010). Normal human bronchial epithelial cells exposed to aerosolized nicotine showed decreased CFTR and BK conductance, impaired CBF, ASL volume, and decreased expression of FOXJ1 and KCNMA1 (Garcia-Arcos et al., 2016).  A concentration-dependent decrease in CBF has been observed after treatment with aldehydes. For example inhibition of cilia ATPase activity was observed after treatment with acetaldehyde, in ciliated bovine bronchial epithelial cells (Sisson et al., 1991). Acrolein, an aldehyde in the gas phase of cigarette smoke, induced ciliostasis at high concentrations (> 1 mM), after 5 min of treatment, and cellular necrosis after 3 hr. However, at lower concentrations (from 0.5‒1 mM), acrolein transiently reduced the CBF to 4 Hz (Romet et al., 1990).

There is no standardized method for measuring CBF. Digital high-speed video imaging with a manual count of CBF in slow motion video play is the most commonly used method for CBF measurement (Kim et al., 2011; Peabody et al., 2018). Photometry and video-microscopy have been used to measure CBF in vitro and ex vivo, including in ciliated bovine bronchial epithelial cells (Allen-Gipson et al., 2011; Sisson et al., 2003; Uzlaner and Priel, 1999), normal human bronchial epithelial cells (Feriani et al., 2017), human nasal epithelial cells (Dimova et al., 2005; Min et al., 1999b), human nasal ciliated epithelium (nasal brushings) (Agius et al., 1998), and mouse tracheal rings (Simet et al., 2010). CBF measurement in vitro generally involves mounting the tissue at the air-liquid interface on a stage followed by microscopic analysis and acquisition of images and/or video recordings of beating cilia. For in vivo and ex vivo measurements, Doppler optical coherence tomography (D-OCT) can also be applied, a mesoscopic non-contact imaging modality that provides high-resolution tomographic images and detects micromotion simultaneously (Jing et al., 2017). D-OCT has been used to quantitatively measure CBF in ex vivo rabbit tracheal cultures (Lemieux et al., 2015).

Age-dependent decreases in CBF have been demonstrated in several species (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013). In a study with 46 healthy subjects with a wide age distribution (mean 42, range 19–81 years), age was found to be negatively associated with airway clearance of inhaled 6-μm Teflon particles (Svartengren et al., 2005). Female hormones, i.e. progesterone and estrogen, have been shown to have direct effect on CBF, i.e., progesterone reduces CBF, 17β-estradiol and progesterone receptor antagonists counteract progesterone effects, but estradiol alone has also been shown to have no effect on CBF. However, the mechanism by which these hormones modulate CBF is yet to be elucidated (Jain et al., 2012; Jia et al., 2011).

UBERON:0000115

UBERON lung epithelium

CL:0005012

CL multi-ciliated epithelial cell

Moderate Mixed

High

All life stages

High High Moderate High High Moderate High

Agius, A. M., L. A. Smallman, and A. L. Pahor (1998). Age, smoking and nasal ciliary beat frequency. Clin. Otolaryngol. Allied Sci. 23, 227-230. Allen-Gipson, D.S., Blackburn, M.R., Schneider, D.J., Zhang, H., Bluitt, D.L., Jarrell, J.C., et al. (2011). Adenosine activation of A(2B) receptor(s) is essential for stimulated epithelial ciliary motility and clearance. Am. J. Physiol. Lung Cell. Mol. Physiol. 301, L171-L180. Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., Devasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589. Cohen, N.A., Zhang, S., Sharp, D.B., Tamashiro, E., Chen, B., Sorscher, E.J., et al. (2009). Cigarette smoke condensate inhibits transepithelial chloride transport and ciliary beat frequency. Laryngoscope 119, 2269-2274. Di Benedetto, G., Manara-Shediac, F.S. and Mehta, A. (1991). Effect of cyclic AMP on ciliary activity of human respiratory epithelium. Eur. Respir. J. 4, 789-795. Dimova, S., Maes, F., Brewster, M.E., Jorissen, M., Noppe, M. and Augustijns, P. (2005). High-speed digital imaging method for ciliary beat frequency measurement. J. Pharmacy Pharmacol 57, 521-526. Feriani, L., Juenet, M., Fowler, C.J., Bruot, N., Chioccioli, M., Holland, S.M., et al. (2017). Assessing the Collective Dynamics of Motile Cilia in Cultures of Human Airway Cells by Multiscale DDM. Biophys. J. 113, 109-119. Garcia-Arcos, I., Geraghty, P., Baumlin, N., Campos, M., Dabo, A.J., Jundi, B., et al. (2016). Chronic electronic cigarette exposure in mice induces features of COPD in a nicotine-dependent manner. Thorax 71, 1119-1129. Gosepath, J., Schaefer, D., Brommer, C., Klimek, L., Amedee, R.G., and Mann, W.J. (2000). Subacute Effects of Ozone Exposure on Cultivated Human Respiratory Mucosa. Am. J. Rhinol. 14, 411-418.  Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B. and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867. Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The Effect of Aging on Nasal Mucociliary Clearance, Beat Frequency, and Ultrastructure of Respiratory Cilia. Am. J. Respir. Crit. Care Med. 163, 983-988. Jain, R., Ray, J.M., Pan, J.-H. and Brody, S.L. (2012). Sex hormone-dependent regulation of cilia beat frequency in airway epithelium. Am. J. Respir. Crit. Care Med. 46, 446-453. Jia, S., Zhang, X., He, D.Z., Segal, M., Berro, A., Gerson, T., et al., 2011. Expression and Function of a Novel Variant of Estrogen Receptor–α36 in Murine Airways. Am. J. Respir. Cell Mol. Biol. 45, 1084-1089. Jiao, J., Wang, H., Lou, W., Jin, S., Fan, E., Li, Y., et al. (2011). Regulation of ciliary beat frequency by the nitric oxide signaling pathway in mouse nasal and tracheal epithelial cells. Exp. Cell Res. 317, 2548-2553. Jing, J.C., Chen, J.J., Chou, L., Wong, B.J.F. and Chen, Z. (2017). Visualization and Detection of Ciliary Beating Pattern and Frequency in the Upper Airway using Phase Resolved Doppler Optical Coherence Tomography. Sci. Rep. 7, 8522-8522. Joki, S. and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea‐pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169. Kim, W., Han, T.H., Kim, H.J., Park, M.Y., Kim, K.S. and Park, R.W. (2011). An Automated Measurement of Ciliary Beating Frequency using a Combined Optical Flow and Peak Detection. J. Healthc. Inform. Res. 17, 111-119. Knoll, M., Shaoulian, R., Magers, T. and Talbot, P. (1995). Ciliary beat frequency of hamster oviducts is decreased in vitro by exposure to solutions of mainstream and sidestream cigarette smoke. Biol. Reprod. 53, 29-37. Kobayashi, K., Tamaoki, J., Sakai, N., Chiyotani, A. and Takizawa, T. (1989). Inhibition of ciliary activity by phorbol esters in rabbit tracheal epithelial cells. Lung 167, 277-284. Kogiso, H., Hosogi, S., Ikeuchi, Y., Tanaka, S., Inui, T., Marunaka, Y., et al. (2018). [Ca(2+) ]i modulation of cAMP-stimulated ciliary beat frequency via PDE1 in airway ciliary cells of mice. Exp. Physiol. 103, 381-390. Lansley, A.B., Sanderson, M.J. and Dirksen, E.R. (1992). Control of the beat cycle of respiratory tract cilia by Ca2+ and cAMP. Am. J. Physiol. 263, L232-242. Lemieux, B.T., Chen, J.J., Jing, J., Chen, Z. and Wong, B.J.F. (2015). Measurement of ciliary beat frequency using Doppler optical coherence tomography. Int. Forum Allergy Rhinol. 5, 1048-1054. Li, D., Shirakami, G., Zhan, X. and Johns, R.A. (2000). Regulation of ciliary beat frequency by the nitric oxide-cyclic guanosine monophosphate signaling pathway in rat airway epithelial cells. Am. J. Respir. Cell Mol. Biol. 23, 175-181. Min, Y.-G., Ohyama, M., Lee, K.S., Rhee, C.-S., Oh, S.H., Sung, M.-W., et al. (1999). Effects of free radicals on ciliary movement in the human nasal epithelial cells. Auris Nasus Larynx 26, 159-163. Paul, P., Johnson, P., Ramaswamy, P., Ramadoss, S., Geetha, B. and Subhashini, A.S. (2013). The Effect of Ageing on Nasal Mucociliary Clearance in Women: A Pilot Study. ISRN Pulmonology 2013, 5. Peabody, J.E., Shei, R.-J., Bermingham, B.M., Phillips, S.E., Turner, B., Rowe, S.M., et al. (2018). Seeing cilia: imaging modalities for ciliary motion and clinical connections. Am. J. Physiol. Lung Cell. Mol. Physiol. 314, L909-L921. Romet, S., Dubreuil, A., Baeza, A., Moreau, A., Schoevaert, D. and Marano, F. (1990). Respiratory tract epithelium in primary culture: Effects of. Toxicol. In Vitro 4, 399-402. Salathe, M., Pratt, M.M. and Wanner, A. (1993). Cyclic AMP-dependent phosphorylation of a 26 kD axonemal protein in ovine cilia isolated from small tissue pieces. Am. J. Respir. Cell Mol. Biol. 9, 306-314. Sanderson, M.J. and Dirksen, E.R. (1989). Mechanosensitive and beta-adrenergic control of the ciliary beat frequency of mammalian respiratory tract cells in culture. Am. Rev. Respir. Dis. 139, 432-440. Schmid, A., Sutto, Z., Nlend, M.-C., Horvath, G., Schmid, N., Buck, J., et al. (2007). Soluble Adenylyl Cyclase Is Localized to Cilia and Contributes to Ciliary Beat Frequency Regulation via Production of cAMP. J. Gen. Physiol. 130, 99-109. Schmid, A., Baumlin, N., Ivonnet, P., Dennis, J.S., Campos, M., Krick, S., et al. (2015). Roflumilast partially reverses smoke-induced mucociliary dysfunction. Respir. Res. 16, 135. Simet, S.M., Sisson, J.H., Pavlik, J.A., Devasure, J.M., Boyer, C., Liu, X., et al. (2010). Long-term cigarette smoke exposure in a mouse model of ciliated epithelial cell function. Am. J. Respir. Cell Mol. Biol. 43, 635-640. Sisson, J.H. (1995). Ethanol stimulates apparent nitric oxide-dependent ciliary beat frequency in bovine airway epithelial cells. Am. J. Physiol. 268, L596-600. Sisson, J.H., May, K. and Wyatt, T.A. (1999). Nitric oxide-dependent ethanol stimulation of ciliary motility is linked to cAMP-dependent protein kinase (PKA) activation in bovine bronchial epithelium. Alcohol Clin. Exp. Res. 23, 1528-1533. Sisson, J.H., Stoner, J., Ammons, B. and Wyatt, T. (2003). All‐digital image capture and whole‐field analysis of ciliary beat frequency. J. Microsc. 211, 103-111. Sisson, J.H., Tuma, D.J. and Rennard, S.I. (1991). Acetaldehyde-mediated cilia dysfunction in bovine bronchial epithelial cells. Am. J. Physiol. 260, L29-36. Svartengren, M., Falk, R. and Philipson, K. (2005). Long-term clearance from small airways decreases with age. Eur. Respir. J. 26, 609-615. Tamaoki, J., Kondo, M. and Takizawa, T. (1989). Effect of cAMP on ciliary function in rabbit tracheal epithelial cells. J. Appl. Physiol. 66, 1035-1039. Uzlaner, N. and Priel, Z. (1999). Interplay between the NO pathway and elevated [Ca(2+)](i) enhances ciliary activity in rabbit trachea. J. Physiol. 516, 179-190. Wang, L.F., White, D.R., Andreoli, S.M., Mulligan, R.M., Discolo, C.M. and Schlosser, R.J. (2012). Cigarette smoke inhibits dynamic ciliary beat frequency in pediatric adenoid explants. Otolaryngol. Head Neck Surg. 146, 659-663. Yaghi, A., Zaman, A., Cox, G. and Dolovich, M.B. (2012). Ciliary beating is depressed in nasal cilia from chronic obstructive pulmonary disease subjects. Respir. Med. 106, 1139-1147. Yang, B., Schlosser, R.J. and Mccaffrey, T.V. (1997). Signal transduction pathways in modulation of ciliary beat frequency by methacholine. Ann. Otol. Rhinol. Laryngol. 106, 230-236. AOPWiki 2021-07-19T10:19:36

2021-09-10T01:38:13

Mucociliary Clearance, Decreased

MCC, Decreased

Individual

In healthy adults, tracheal mucus movement varies from 4 to >20 mm/min (Stannard and O'Callaghan, 2006), whereas mucociliary clearance (MCC) in the small airways is slower due to the lower number of ciliated cells (fewer cilia) and their shorter length (Foster et al., 1980; Iravani, 1969; Wanner et al., 1996). Since optimal MCC is dependent in multiple factors, including cilia number and structure as well as ASL and mucus properties, any disturbances of these can lead to impaired MCC. While high humidity or infection can enhance MCC, long-term exposure to noxious substances (e.g. cigarette smoke) lead to decreased mucus clearance from the airways. In most instances this is reflected by decreased mucus transport rates or velocities.

In humans, MCC has been assessed traditionally following inhalation of radio-labeled particles such as 99Tcm-labeled polystyrene particles, resin particles or serum albumin and following their clearance at regular intervals by radioimaging using gamma cameras (Agnew et al., 1986; Kärjä et al., 1982). Taking into account inhalation volumes and flow rates, lung airflow, particle deposition and retention, clearance rates can be calculated and effects of e.g. drugs on MCC can be examined. Alternatively, since MCC occurs at a similar rate in the nose to that in trachea and bronchi (Andersen and Proctor, 1983; Rutland and Cole, 1981) and for ease of use, measurements of MCC can be restricted to that of nasal MCC only. Probably one of the simplest methods is the saccharin transit test (STT). For this test, a small particle of saccharin is placed behind the anterior end of the inferior turbinate. The saccharin will be transported by mucociliary action toward the nasopharynx, where its sweet taste is perceived. When MCC is impaired, saccharin transit times will increase, with a 10- to 20-minute delay being considered a clinical sign of decreased MCC. Using the same principle, the test can also be performed or complemented with dyes such as indigo carmine or methylene blue (Deborah and Prathibha, 2014). In experimental animals, MCC has been evaluated by gamma-scintigraphy (Greiff et al., 1990; Hua et al., 2010; Read et al., 1992), fluorescence videography/fluoroscopy (in explanted tracheas etc.) (Grubb et al., 2016; Rogers  et al., 2018), or by 3D-SPECT (Ortiz Belda et al., 2016). Direct observation of particle movement across airway epithelia to determine mucus velocity or transport rates by using a fiberoptic bronchoscope may be helpful when working in larger animals such as dogs (King, 1998). In vitro, freshly excised frog palate preparations have been used to assess cilia function and mucociliary transport by videomicroscopy (Macchione et al., 1995; Macchione et al., 1999; Trindade et al., 2007). Murine and human nasal, bronchial and small airway epithelial models grown at the air-liquid interface are also suitable in vitro test systems for determining mucus transport by tracing inert particle movement with a set-up similar to that used for assessing CBF (Benam et al., 2018; Fliegauf et al., 2013; Knowles and Boucher, 2002; Sears et al., 2015).

High Mixed

High

All life stages

High Moderate Moderate Moderate Moderate Moderate Moderate

Agnew, J., Sutton, P., Pavia, D. and Clarke, S. (1986). Radioaerosol assessment of mucociliary clearance: towards definition of a normal range. Brit. J. Radiol. 59, 147-151. Allegra, L., Moavero, N., and Rampoldi, C. (1991). Ozone-induced impairment of mucociliary transport and its prevention with N-acetylcysteine. Am. J. Med. 91, S67-S71. Andersen, I. and Proctor, D. (1983). Measurement of nasal mucociliary clearance. Eur. J. Respir. Dis. Suppl. 127, 37-40. Baby, M.K., Muthu, P.K., Johnson, P., and Kannan, S. (2014). Effect of cigarette smoking on nasal mucociliary clearance: A comparative analysis using saccharin test. Lung India 31, 39-42.  Benam, K.H., Vladar, E.K., Janssen, W.J. and Evans, C.M. (2018). Mucociliary defense: emerging cellular, molecular, and animal models. Ann. Am. Thorac. Soc. 15, S210-S215. Deborah, S. and Prathibha, K., 2014. Measurement of nasal mucociliary clearance. Clin. Res. Pulmonol. 2, 1019. Dülger, S., Akdeniz, Ö., Solmaz, F., Şengören Dikiş, Ö., and Yildiz, T. (2018). Evaluation of nasal mucociliary clearance using saccharin test in smokers: A prospective study. Clin. Respir. J. 12, 1706-1710.  Fliegauf, M., Sonnen, A.F.P., Kremer, B. and Henneke, P. (2013). Mucociliary Clearance Defects in a Murine In Vitro Model of Pneumococcal Airway Infection. PloS ONE 8, e59925. Fló-Neyret, C., Lorenzi-Filho, G., Macchione, M., Garcia, M.L.B. and Saldiva, P.H.N. (2001). Effects of formaldehyde on the frog's mucociliary epithelium as a surrogate to evaluate air pollution effects on the respiratory epithelium. Braz. J. Med. Biol. Res. 34, 639-643. Foster, W., Langenback, E. and Bergofsky, E. (1980). Measurement of tracheal and bronchial mucus velocities in man: relation to lung clearance. J. Appl. Physiol. 48, 965-971. Greiff, L., Wollmer, P., Erjefält, I., Pipkorn, U. and Persson, C. (1990). Clearance of 99mTc DTPA from guinea pig nasal, tracheobronchial, and bronchoalveolar airways. Thorax 45, 841-845. Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B. and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867. Habesoglu, M., Demir, K., Yumusakhuylu, A.C., Sahin Yilmaz, A., and Oysu, C. (2012). Does passive smoking have an effect on nasal mucociliary clearance? Otolaryngol Head Neck Surg. 147, 152-156. Hua, X., Zeman, K.L., Zhou, B., Hua, Q., Senior, B.A., Tilley, S.L., et al. (2010). Noninvasive real-time measurement of nasal mucociliary clearance in mice by pinhole gamma scintigraphy. J. Appl. Physiol. 108, 189-196. Iravani, J. (1969). Zum Mechanismus der Ortsabhängigkeit der Flimmeraktivität im Bronchialbaum/Location-Dependent Activity of the Ciliary Movement in the Bronchial Tree and its Possible Mechanism. In: Habermann E. et al. (eds) Naunyn Schmiedebergs Archiv für Pharmakologie. Springer, Berlin, Heidelberg. Kakinoki Y, Ohashi Y, Tanaka A, Washio Y, Yamada K, Nakai Y, Morimoto K. (1998). Nitrogen dioxide compromises defence functions of the airway epithelium. Acta Oto-Laryngol. 118, 221-226. Kärjä, J., Nuutinen, J. and Karjalainen, P. (1982). Radioisotopic Method for Measurement of Nasal Mucociliary Activity. Arch. Otolaryngol. 108, 99-101. King, M. (1998). Experimental models for studying mucociliary clearance. Eur. Respir. J. 11, 222-228. Knorst, M.M., Kienast, K., Riechelmann, H., Müller-Quernheim, J. and Ferlinz, R. (1994). Effect of sulfur dioxide on mucociliary activity and ciliary beat frequency in guinea pig trachea. Int. Arch. Occup. Environm. Health 65, 325-328. Knowles, M.R. and Boucher, R.C. (2002). Mucus clearance as a primary innate defense mechanism for mammalian airways. J. Clin. Invest. 109, 571-577. Macchione, M., Guimarães, E., Saldiva, P. and Lorenzi-Filho, G. (1995). Methods for studying respiratory mucus and mucus clearance. Braz. J. Med. Biol Res. 28, 1347. Macchione, M., Oliveira, A.P., Gallafrio, C.T., Muchão, F.P., Obara, M.T., Guimarães, E.T., et al. (1999). Acute effects of inhalable particles on the frog palate mucociliary epithelium. Environm. Health Persp. 107, 829-833. Morgan, K., Patterson, D. and Gross, E. (1986). Responses of the nasal mucociliary apparatus of F-344 rats to formaldehyde gas. Toxicol. Appl. Pharmacol. 82, 1-13. Morgan, K.T., Patterson, D.L. and Gross, E.A. (1984). Frog palate mucociliary apparatus: structure, function, and response to formaldehyde gas. Fund. Appl. Toxicol. 4, 58-68. Ortiz Belda, J.L., Ortiz, A., Milara Payá, J., Armengot Carceller, M., Sanz García, C., Compañ Quilis, D., et al. (2016). Evaluation of Mucociliary Clearance by Three Dimension Micro-CT-SPECT in Guinea Pig: Role of Bitter Taste Agonists. Plos ONE 11, e0164399. Pagliuca, G., Rosato, C., Martellucci, S., De Vincentiis, M., Greco, A., Fusconi, M., et al. (2015). Cytologic and functional alterations of nasal mucosa in smokers: temporary or permanent damage? Otolaryngol Head Neck Surg 152, 740-745. Proença, M., Xavier, R.F., Ramos, D., Cavalheri, V., Pitta, F., and Ramos, E.C. (2011). Immediate and short term effects of smoking on nasal mucociliary clearance in smokers. Revista Portuguesa de Pneumologia (English Edition) 17), 172-176. Read, R.C., Roberts, P., Munro, N., Rutman, A., Hastie, A., Shryock, T., et al. (1992). Effect of Pseudomonas aeruginosa rhamnolipids on mucociliary transport and ciliary beating. J. Appl. Physiol. 72, 2271-2277. Rogers, T.D., Ostrowski, L.E., Livraghi-Butrico, A., Button, B. and Grubb, B.R., 2018. Mucociliary clearance in mice measured by tracking trans-tracheal fluorescence of nasally aerosolized beads. Sci. Rep. 8, 1-12. Rutland, J. and Cole, P.J. (1981). Nasal mucociliary clearance and ciliary beat frequency in cystic fibrosis compared with sinusitis and bronchiectasis. Thorax 36, 654-658. Sears, P.R., Yin, W.-N. and Ostrowski, L.E. (2015). Continuous mucociliary transport by primary human airway epithelial cells in vitro. Am. J. Physiol. Lung Cell. Mol. Physiol. 309, L99-L108. Solak, I., Marakoglu, K., Pekgor, S., Kargin, N.C., Alataş, N., and Eryilmaz, M.A. (2018). Nasal mucociliary activity changes in smokers. Konuralp Med. J. 10, 269-275. Stannard, W. and O'callaghan, C. (2006). Ciliary function and the role of cilia in clearance. J. Aerosol Med. 19, 110-115. Trindade, S.H.K., De Mello Júnior, J.F., De Godoy Mion, O., Lorenzi-Filho, G., Macchione, M., Guimarães, E.T., et al. (2007). Methods for Studying Mucociliary Transport. Braz. J. Otorhinolaryngol. 73, 704-712. Wanner, A., Salathe, M. and O'riordan, T.G. (1996). Mucociliary clearance in the airways. Am. J. Respir. Crit. Care Med. 154, 1868-1902. Xavier, R.F., Ramos, D., Ito, J.T., Rodrigues, F.M., Bertolini, G.N., Macchione, M., et al. (2013). Effects of cigarette smoking intensity on the mucociliary clearance of active smokers. Respiration 86, 479-485.  Yadav, J., and Kaushik, G. (2014). K Ranga R. Passive smoking affects nasal mucociliary clearance. J. Indian Acad. Clin. Med. 15, 96-99. Yeates, D.B., Katwala, S.P., Daugird, J., Daza, A.V. and Wong, L.B. (1997). Excitatory and inhibitory neural regulation of tracheal ciliary beat frequency (CBF) activated by ammonia vapour and SO2. Ann. Occup. Hyg. 41, 736-744. AOPWiki 2021-07-19T10:21:38

2021-09-10T07:19:28

Decrease, Lung function

Decreased lung function

Individual

Lung function is a clinical term referring to the physiological functioning of the lungs, most often in association with the tests used to assess it. Lung function loss can be caused by acute or chronic exposure to airborne toxicants or by an intrinsic disease of the respiratory system.  Although signs of cellular injury are typically exhibited first in the nose and larynx, alveolar-capillary barrier breakdown may ultimately arise and result in local edema (Miller and Chang, 2003). Clinically, bronchoconstriction and hypoxia are seen in the acute phase, with affected subjects exhibiting shortness of breath (dyspnea) and low blood oxygen saturation, and with reduced lung function indices of airflow, lung volume and gas exchange (Hert and Albert, 1994; and How it is Measured or Detected;). When alveolar damage is extensive, the reduced lung function can develop into acute respiratory distress syndrome (ARDS). This severe compromise of lung function is reflected by decreased gas exchange indices (PaO2/FIO2 ≤200 mmHg, due to hypoxemia and impaired excretion of carbon dioxide), increased pulmonary dead space and decreased respiratory compliance (Matthay et al., 2019). Acute inhalation exposures to chemical irritants such as ammonia, hydrogen chloride, nitrogen oxides and ozone typically cause local edema that manifests as dyspnea and hypoxia. In cases where a breakdown of the alveolar capillary function ensues, ARDS develops. ARDS has a particularly high risk of mortality, estimated to be 30-40% (Gorguner and Akgun, 2010; Matthay et al., 2018; Reilly et al., 2019). Lung function decrease due to reduction in lung volume is seen in pulmonary fibrosis, which can be linked to chronic exposures to e.g. silica, asbestos, metals, agricultural and animal dusts (Meltzer and Noble, 2008; Cheresh et al., 2013; Cosgrove, 2015; Trethewey and Walters, 2018). Additionally. decreased lung function occurs in pleural disease, chest wall and neuromuscular disorders, because of obesity and following pneumectomy (Moore, 2012). Decreased lung function can also be a result of narrowing of the airways by inflammation and mucus plugging resulting in airflow limitation. Decreased lung function is a feature of obstructive pulmonary diseases (e.g. asthma, COPD) and linked to a multitude of causes, including chronic exposure to cigarette smoke, dust, metals, organic solvents, asbestos, pathogens or genetic factors.

Pulmonary function tests are a group of tests that evaluate several parameters indicative of lung size, air flow and gas exchange. Decreased lung function can manifest in different ways, and individual circumstances, including potential exposure scenarios, determine which test is used. The section outlines the tests used to evaluate lung function in humans (https://www.nhlbi.nih.gov/health-topics/pulmonary-function-tests, accessed 22 March 2021) and in experimental animals. Lung function tests used to evaluate human lung function The most common (“gold standard”) lung function test in human subjects is spirometry. Spirometry results are primarily used for diagnostic purposes, e.g. to discriminate between obstructive and restrictive lung diseases, and for determining the degree of lung function impairment. Specific criteria for spirometry tests have been outlined in the American Thoracic Society (ATS) and the European Respiratory Society (ERS) Task Force guidelines (Graham et al., 2019). These guidelines consist of detailed recommendations for the preparation and conduct of the test, instruction of the person tested, as well as indications and contraindications, and are complemented by additional guidance documents on how to interpret and report the test results (Pellegrino et al., 2005; Culver et al., 2017). Spirometry measures several different parameters during forceful exhalation, including: <ul> <li style="text-align:justify">Forced expiratory volume in 1 s (FEV1), the maximum volume of air that can forcibly be exhaled during the first second following maximal inhalation</li> <li style="text-align:justify">Forced vital capacity (FVC), the maximum volume of air that can forcibly be exhaled following maximal inhalation </li> <li style="text-align:justify">Vital capacity (VC), the maximum volume of air that can be exhaled when exhaling as fast as possible</li> <li style="text-align:justify">FEV1/FVC ratio</li> <li style="text-align:justify">Peak expiratory flow (PEF), the maximal flow that can be exhaled when exhaling at a steady rate</li> <li style="text-align:justify">Forced expiratory flow, also known as mid-expiratory flow; the rates at 25%, 50% and 75% FVC are given</li> <li style="text-align:justify">Inspiratory vital capacity (IVC), the maximum volume of air that can be inhaled after a full expiration</li> </ul> A reduced FEV1, with normal or reduced VC, normal or reduced FVC, and a reduced FEV1/FVC ratio are indices of airflow limitation, i.e., airway obstruction as seen in COPD (Moore, 2012). In contrast, airway restriction is demonstrated by a reduction in FVC, normal or increased FEV1/FVC ratio, a normal spirometry trace and potentially a high PEF (Moore, 2012). Lung capacity or lung volumes can be measured using one of three basic techniques: 1) plethysmography, 2) nitrogen washout, or 3) helium dilution. Plethysmography consists of a series of sequential measurements in a body plethysmograph, starting with the measurement of functional residual capacity (FRC), the volume of gas present in the lung at end-expiration during tidal breathing. Once the FRC is known, expiratory reserve volume (ERV; the volume of gas that can be maximally exhaled from the end-expiratory level during tidal breathing, i.e., the FRC), vital capacity (VC; the volume change at the mouth between the positions of full inspiration and complete expiration), and inspiratory capacity (IC; the maximum volume of air that can be inhaled from FRC) are determined, and total lung capacity (TLC; the volume of gas in the lungs after maximal inspiration, or the sum of all volume compartments) and residual volume (RV; the volume of gas remaining in the lung after maximal exhalation) are calculated (Weinstock and McCannon, 2017). The other two techniques used to measure lung volumes—helium dilution and nitrogen washout—are based on the principle of conservation of mass: [initial gas concentration] x [initial volume of the system] = [final gas concentration] x [final volume of the system]. The nitrogen washout method is based on the fact that nitrogen is present in the air, at a relatively constant amount. The subject is given 100% oxygen to breathe, and the expired gas, which contains nitrogen in the lung at the beginning of the test, is collected. When no more nitrogen is noted in the expirate, the volume of air expired and the entire amount of nitrogen in that volume are measured, and the initial volume of the system (FRC) can be calculated. In the helium dilution method, a known volume and concentration of helium is inhaled by the subject. Helium, an inert gas that is not absorbed significantly from the lungs, is diluted in proportion to the lung volume to which it is added. The final concentration of helium is then measured and FRC calculated (Weinstock and McCannon, 2017). Measurements of lung volumes in humans are technically more challenging than spirometry. However, they complement spirometry (which cannot determine lung volumes) and may be a preferred means of lung function assessment when subject compliance cannot be reasonably expected (e.g. in pediatric subjects) or where forced expiratory maneuvers are not possible (e.g. in patients with advanced pulmonary fibrosis). There are recommended standards for lung volume measurements and their interpretation in clinical practice, issued by the ATS/ERS Task Force (Wanger et al., 2005; Criée et al., 2011). Finally, indices of gas exchange across the alveolar-capillary barrier are tested by diffusion capacity of carbon monoxide (DLCO) studies (also referred to as transfer capacity of carbon monoxide, TLCO). The principle of the test is the increased affinity of hemoglobin to preferentially bind carbon monoxide over oxygen (Weinstock and McCannon, 2017). Complementary to spirometry and lung volume measurements, DLCO provides information about the lung surface area available for gas diffusion. Therefore, it is sensitive to any structural changes affecting the alveoli, such as those accompanying emphysema, pulmonary fibrosis, pulmonary edema, and ARDS. Recommendations for the standardization of the test and its evaluation have been outlined by the ATS/ERS Task Force (Graham et al., 2017). An isolated reduction in DLCO with normal spirometry and in absence of anemia suggests an injury to the alveolar-capillary barrier, as for example seen in the presence of pulmonary emboli or in patients with pulmonary hypertension (Weinstock and McCannon, 2017; Lettieri et al., 2006; Seeger et al., 2013). Reduced DLCO together with airflow obstruction (i.e., reduced FEV1) indicates lung parenchymal damage and is commonly observed in smokers and in COPD patients (Matheson et al., 2007; Harvey et al., 2016), whereas reduced DLCO with airflow restriction is seen in patients with interstitial lung diseases (Dias et al., 2014; Kandhare et al., 2016). Lung function tests used to evaluate experimental animal lung function Because spirometry requires active participation and compliance of the subject, it is not commonly used in animal studies. However, specialized equipment such as the flexiVent system (SCIREQ®) are available for measuring FEV, FVC and PEF in anesthetized and tracheotomized small laboratory animals. Other techniques such as plethysmography or forced oscillation are increasingly preferred for lung function assessment in small laboratory animals (McGovern et al., 2013; Bates, 2017). In small laboratory animals, plethysmography can be used to determine respiratory physiology parameters (minute volume, respiratory rate, time of pause and time of break), lung volume and airway resistance of conscious animals. Both whole body and head-out plethysmography can be applied, although there is a preference for the latter in the context of inhalation toxicity studies, because of its higher accuracy and reliability (OECD, 2018a; Hoymann, 2012). Gas diffusion tests are not frequently performed in animals, because reproducible samplings of alveolar gas are difficult and technically challenging (Reinhard et al., 2002; Fallica et al., 2011). Modifications to the procedure employed in humans have, however, open possibilities to obtain a human-equivalent DLCO measure or the diffusion factor for carbon monoxide (DFCO)—a variable closely related to DLCO, which can inform on potential structural changes in the lungs that have an effect on gas exchange indices (Takezawa et al., 1980; Dalbey et al., 1987; Fallica et al., 2011; Limjunyawong et al., 2015).

Pulmonary function tests reflect the physiological working of the lungs. Therefore, the AO is applicable to a variety of species, including (but not limited to) rodents, rabbits, pigs, cats, dogs, horses and humans, independent of life stage and gender.

High Mixed

High

Adult

High

<div style="text-align:justify"> Ackermann-Liebrich, U., Leuenberger, P., Schwartz, J., Schindler, C., Monn, C., Bolognini, G., et al. (1997). Lung function and long term exposure to air pollutants in Switzerland. Study on Air Pollution and Lung Diseases in Adults (SAPALDIA) Team. Am. J. Resp. Crit. Care Med. 155, 122-129.  Adam, M., Schikowski, T., Carsin, A.E., Cai, Y., Jacquemin, B., Sanchez, M., et al. (2015). Adult lung function and long-term air pollution exposure. ESCAPE: a multicentre cohort study and meta-analysis. Eur. Resp. J. 45, 38-50. Andersen, M.H.G., Frederiksen, M., Saber, A.T., Wils, R.S., Fonseca, A.S., Koponen, I.K., et al. (2019). Health effects of exposure to diesel exhaust in diesel-powered trains. Part. Fibre Toxicol. 16, 21. Ashley, F., Kannel, W.B., Sorlie, P.D., and Masson, R. (1975). Pulmonary function: relation to aging, cigarette habit, and mortality: the Framingham Study. Ann. Int. Med. 82, 739-745. Bates, J.H.T. (2017). CO Bergstra, A.D., Brunekreef, B., and Burdorf, A. (2018). The effect of industry-related air pollution on lung function and respiratory symptoms in school children. Environm. Health 17, 30.  Baumgartner, K.B., Samet, J.M., Coultas, D.B., Stidley, C.A., Hunt, W.C., Colby, T.V., and J.A. Waldron (2000). Occupational and environmental risk factors for idiopathic pulmonary fibrosis: a multicenter case-control study. Collaborating Centers. Am. J. Epidemiol. 152, 307-315. Broekema, M., ten Hacken, N.H., Volbeda, F., Lodewijk, M.E., Hylkema, M.N., Postma, D.S., et al. (2009). Airway epithelial changes in smokers but not in ex-smokers with asthma. Am. J. Resp. Crit. Care Med. 180, 1170-1178. Celli, B. R. (2010). Predictors of mortality in COPD. Respir. Med. 104, 773-779. Cheresh, P., Kim, S.J., Tulasiram, S., and D.W. Kamp (2013). Oxidative stress and pulmonary fibrosis. Biochim. Biophys. Acta, 1832, 1028–1040. Cosgrove, M.P. (2015). Pulmonary fibrosis and exposure to steel welding fume. Occup. Med. 65, 706-712. Criée, C.P., Sorichter, S., Smith, H.J., Kardos, P., Merget, R., Heise, D., Berdel, D., Köhler, D., Magnussen, H., Marek, W. and H. Mitfessel (2011). Body plethysmography–its principles and clinical use. Respir. Med. 105, 959-971. Dalbey, W., Henry, M., Holmberg, R., Moneyhun, J., Schmoyer, R. and S. Lock (1987). Role of exposure parameters in toxicity of aerosolized diesel fuel in the rat. J. Appl. Toxicol. 7, 265-275. Dias, O.M., Baldi, B.G., Costa, A.N., C.R. Carvalho (2014). Combined pulmonary fibrosis and emphysema: an increasingly recognized condition. J. Bras. Pneumol. 40, 304-312.  Fallica, J., Das, S., Horton, M., and W. Mitzner (2011). Application of carbon monoxide diffusing capacity in the mouse lung. J. Appl. Physiol. 110, 1455–1459. Forbes, L.J., Kapetanakis, V., Rudnicka, A.R., Cook, D.G., Bush, T., Stedman, J.R., et al. (2009). Chronic exposure to outdoor air pollution and lung function in adults. Thorax 64, 657-663. 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Progression to COPD in smokers with normal spirometry/low DLCO using different methods to determine normal levels. Eur. Respir. J. 47, 1888-1889. Hert, R. and R.K. Albert (1994). Sequelae of the adult respiratory distress syndrome. Thorax 49, 8-13. Hoymann, H.G. (2012). Lung function measurements in rodents in safety pharmacology studies. Front. Pharmacol. 3, 156. Johnson, J. D., and W. M. Theurer (2014). A stepwise approach to the interpretation of pulmonary function tests. Am. Fam. Phys. 89, 359-366. Kandhare, A.D., Mukherjee, A., Ghosh, P. and S.L. Bodhankar (2016). Efficacy of antioxidant in idiopathic pulmonary fibrosis: A systematic review and meta-analysis. EXCLI J. 15, 636. Kim, C.S., Alexis, N.E., Rappold, A.G., Kehrl, H., Hazucha, M.J., Lay, J.C., et al. (2011). Lung function and inflammatory responses in healthy young adults exposed to 0.06 ppm ozone for 6.6 hours. Am. J. Respir. Crit. Care Med. 183, 1215-1221. Kuperman, A.S., and Riker, J.B. (1973). The variable effect of smoking on pulmonary function. Chest 63, 655-660.  Lee, H. M., Liu, M. A., Barrett-Connor, E., and N. D. Wong (2014). Association of Lung Function with Coronary Heart Disease and Cardiovascular Disease Outcomes in Elderly: The Rancho Bernardo Study. Respir. Med. 108, 1779–1785. Lettieri, C.J., Nathan, S.D., Barnett, S.D., Ahmad, S. and A.F. Shorr (2006). Prevalence and outcomes of pulmonary arterial hypertension in advanced idiopathic pulmonary fibrosis. Chest 129, 746-752. Limjunyawong, N., Fallica, J., Ramakrishnan, A., Datta, K., Gabrielson, M., Horton, M., and W. Mitzner (2015). Phenotyping mouse pulmonary function in vivo with the lung diffusing capacity. JoVE 95, e52216. Matheson, M.C., Raven, J., Johns, D.P., Abramson, M.J. and E.H. Walters (2007). Associations between reduced diffusing capacity and airflow obstruction in community-based subjects. Respir. Med. 101, 1730-1737. Matthay, M.A., Zemans, R.L., Zimmerman, G.A., Arabi, Y.M., Beitler, J.R., Mercat, A., Herridge, M., Randolph, A.G. and C.S. Calfee (2019). Acute respiratory distress syndrome. Nature Reviews Disease Primers 5, 1-22. McGovern, T.K., Robichaud, A., Fereydoonzad, L., Schuessler, T.F., and J.G. Martin (2013) Evaluation of respiratory system mechanics in mice using the forced oscillation technique. JoVE 75, e50172. Meltzer, E.B., and P.W. Noble (2008). Idiopathic pulmonary fibrosis. Orphanet J. Rare Dis. 3, 8. Miller, K. and A. Chang (2003). Acute inhalation injury. Emerg. Med. Clin. N. Am. 21, 533-557. Miller, M.R., Crapo, R., Hankinson, J., Brusasco, V., Burgos, F., Casaburi, R., Coates, A., Enright, P., van der Grinten, C.M., and P. Gustafsson (2005a). General considerations for lung function testing. Eur. Respir. J. 26, 153-161. Miller, M.R., Hankinson, J., Brusasco, V., Burgos, F., Casaburi, R., Coates, A., Crapo, R., Enright, P., van der Grinten, C., and P. Gustafsson (2005b). Standardisation of spirometry. Eur. Respir. J. 26, 319-338. Moore, V.C. (2012). Spirometry: step by step. Breathe 8, 232-240. OECD (2018a). OECD Guidance Document on Inhalation Toxicity Studies, GD 39. OECD (2018b), Test No. 412: Subacute Inhalation Toxicity: 28-Day Study, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris, https://doi.org/10.1787/9789264070783-en. OECD (2018), Test No. 413: Subchronic Inhalation Toxicity: 90-day Study, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris, https://doi.org/10.1787/9789264070806-en. Park, Y., Ahn, C., and T.H. Kim (2021) Occupational and environmental risk factors of idiopathic pulmonary fibrosis: a systematic review and meta-analyses. Sci. Rep. 11, 4318. Prada-Dacasa, P., Urpi, A., Sánchez-Benito, L., Bianchi, P., A. Quintana (2020). Measuring Breathing Patterns in Mice Using Whole-body Plethysmography. Bio. Protoc. 10, e3741. Pelkonen, M., Notkola, I.-L., Nissinen, A., Tukiainen, H., and Koskela, H. (2006). Thirty-year cumulative incidence of chronic bronchitis and COPD in relation to 30-year pulmonary function and 40-year mortality: a follow-up in middle-aged rural men. Chest 130, 1129-1137. Pellegrino, R., Viegi, G., Brusasco, V., Crapo, R., Burgos, F., Casaburi, R., Coates, A., van der Grinten, C., Gustafsson, P., and J. Hankinson (2005). Interpretative strategies for lung function tests. Eur. Respir. J. 26, 948-968. Raghu, G., Remy-Jardin, M., Myers, J.L., Richeldi, L., Ryerson, C.J., Lederer, D.J., Behr, J., Cottin, V., Danoff, S.K., Morell, F., and K.R. Flaherty (2018). Diagnosis of idiopathic pulmonary fibrosis. An official ATS/ERS/JRS/ALAT clinical practice guideline. Am. J. Respir. Crit. Care Med. 198, e44-e68. Reilly, J.P., Zhao, Z., Shashaty, M.G., Koyama, T., Christie, J.D., Lanken, P.N., Wang, C., Balmes, J.R., Matthay, M.A., Calfee, C.S. and L.B. Ware (2019). Low to moderate air pollutant exposure and acute respiratory distress syndrome after severe trauma. Am. J. Respir. Crit. Care Med. 199, 62-70. Reinhard. C., Eder, G., Fuchs, H., Ziesenis, A., Heyder, J. and H. Schulz H (2002). Inbred strain variation in lung function. Mamm. Genome 13, 429-437. RP: Measurement of lung function in small animals. J. Appl. Physiol. 123, 1039-1046. Schikowski, T., Sugiri, D., Ranft, U., Gehring, U., Heinrich, J., Wichmann, H.E., et al. (2005). Long-term air pollution exposure and living close to busy roads are associated with COPD in women. Respir. Res. 6, 152.  Seeger, W., Adir, Y., Barberà, J.A., Champion, H., Coghlan, J.G., Cottin, V., De Marco, T., Galiè, N., Ghio, S., Gibbs, S. and F.J. Martinez (2013). Pulmonary hypertension in chronic lung diseases. J. Am. Coll. Cardiol. 62 Suppl. 25, D109-D116. Sin, D. D., Wu, L., and S. P. Man (2005). The relationship between reduced lung function and cardiovascular mortality: a population-based study and a systematic review of the literature. Chest 127, 1952-1959. Takezawa, J., Miller, F.J. and J.J. O'Neil (1980). Single-breath diffusing capacity and lung volumes in small laboratory mammals. J. Appl. Physiol. 48, 1052-1059. Tantisuwat, A., and Thaveeratitham, P. (2014). Effects of smoking on chest expansion, lung function, and respiratory muscle strength of youths. J. Phys. Ther. Sci. 26, 167-170.  Trethewey, S. P., and G. I. Walters (2018). The Role of Occupational and Environmental Exposures in the Pathogenesis of Idiopathic Pulmonary Fibrosis: A Narrative Literature Review. Medicina (Kaunas, Lithuania) 54, 108. Tsui, H.-C., Chen, C.-H., Wu, Y.-H., Chiang, H.-C., Chen, B.-Y., and Guo, Y.L. (2018). Lifetime exposure to particulate air pollutants is negatively associated with lung function in non-asthmatic children. Environ. Poll. 236, 953-961.  Vestbo, J., Anderson, W., Coxson, H.O., Crim, C., Dawber, F., Edwards, L., Hagan, G., Knobil, K., Lomas, D.A., MacNee, W. and E.K. Silverman (2008). Evaluation of COPD longitudinally to identify predictive surrogate end-points (ECLIPSE). Eur. Respir. J. 31, 869-73. Wanger, J., Clausen, J.L., Coates, A., Pedersen, O.F., Brusasco, V., Burgos, F., Casaburi, R., Crapo, R., Enright, P., Van Der Grinten, C.P.M. and P. Gustafsson (2005). Standardisation of the measurement of lung volumes. Eur. Respir. J. 26, 511-522. Weinstock, T, and J. McCannon (2017). Pulmonary Medicine. Pulmonary Function Testing. https://www.pulmonologyadvisor.com/home/decision-support-in-medicine/pulmonary-medicine/pulmonary-function-testing/ (accessed 22 March 2021). Decision Support in Medicine, LLC. Wise, R. A. (2006). The value of forced expiratory volume in 1 second decline in the assessment of chronic obstructive pulmonary disease progression. Am. J. Med. 119, 4-11. Zhang, L.P., Zhang, X., Duan, H.W., Meng, T., Niu, Y., Huang, C.F., et al. (2017). Long-term exposure to diesel engine exhaust induced lung function decline in a cross sectional study. Ind.l Health 55, 13-26. </div> AOPWiki 2016-11-29T18:41:31

2021-09-08T04:54:28

<upstream-id>bd771a49-63b3-4dd8-bfd6-5d62ed89b901</upstream-id> <downstream-id>381aa3f3-9bed-41f2-afe5-10ce883fe336</downstream-id> Oxidative stress (such as that caused by cigarette smoke exposure or irradiation) leads to decreased forkhead box J1 (FOXJ1) gene and protein expression, as well as to decreased FOXJ1 target gene expression (Brekman et al., 2014; Garcia-Arcos et al., 2016; Ishikawa and Ito, 2017; Milara et al., 2012; Valencia-Gattas et al., 2016). FOXJ1 is a key factor of multiple motile cilia assembly in the respiratory airways (Zhou and Roy, 2015). Thus oxidative stress blocks the multiple ciliogenesis program in the airway epithelium.

Cigarette smoke-induced oxidative stress downregulates FOXJ1 expression at both the gene and protein levels in human lung cells in vitro (Milara et al., 2012; Brekman et al., 2014; Valencia-Gattas et al., 2016; Ishikawa and Ito, 2017). Oxidative stress induced by human respiratory syncytial virus reduces FOXJ1 mRNA levels, which can be restored by treatment with antioxidants or the phosphodiesterase 4 inhibitor roflumilast N-oxide (Akaike et al., 1990; Geiler et al., 2010; Mata et al., 2012). In mice, thoracic irradiation results in free radical generation and subsequent reduction in FOXJ1 mRNA expression (Bernard et al., 2012). Many genes that are transcriptionally regulated by FOXJ1 are also downregulated following exposure to cigarette smoke, which implies a reduction in FOXJ1 transcriptional activity (Brekman et al., 2014).

The negative association between cigarette smoke exposure and FOXJ1 levels in airways was shown in multiple studies and can be estimated as a strong linkage. Yet, the notion that oxidative stress as a result of cigarette smoke exposure is leading to decreased FOXJ1 levels is not well demonstrated. As a complex mixture of thousands of chemicals, cigarette smoke exposure could lead to reduced FOXJ1 levels via different routes. Indirect evidence, such as antioxidant molecules that restore cigarette smoke exposure-reduced FOXJ1 levels, as well evidences from other oxidative stress generating insults that decrease FOXJ1 levels add confidence to this KER. However, studies showing a link between oxidative stress generating agents and reduced FOXJ1 levels are scarce. Collectively, the empirical evidence and uncertainties of the linkage imply a moderate ranking for the KER.

Whole cigarette smoke exposed normal human bronchial epithelial cells (HBECs) had significantly lower FoxJ1 protein levels than air-exposed controls (by immunofluorescence). In addition, FOXJ1 mRNA levels were reduced in whole smoke-exposed differentiating and differentiated HBECs (by TaqMan quantitative RT-PCR)(Valencia-Gattas et al., 2016). Treatment with cigarette smoke extract (CSE) significantly down-regulated FOXJ1 mRNA and protein levels in differentiating human airway basal cells in air–liquid interface (ALI) cultures (by TaqMan qRT-PCR and Western blot analysis) (Brekman et al., 2014). Treatment with cigarette smoke extract (CSE) reduced FOXJ1 gene and protein expression in differentiated HBECs through an IL13-mediated mechanism (by RT-qPCR and Western blot analysis) (Milara et al., 2012). Treatment with roflumilast N-oxide (which reduced intracellular reactive oxygen species levels) prevented FOXJ1 loss in CSE-treated cells (Milara et al., 2012). Exposure of 3D co-cultures of HBECs and fibroblasts to whole cigarette smoke decreased FOXJ1 gene expression in a concentration-dependent manner (by TaqMan quantitative RT-PCR) (Ishikawa and Ito, 2017). Human respiratory syncytial virus (RSV) infections involve reactive oxygen intermediates (ROIs) that cause cellular damage (Akaike et al., 1990; Mata et al., 2012). Treatment with the free radical scavenger N-acetylcysteine (NAC) reduced the RSV inflammatory response (Geiler et al., 2010). RSV infection reduced FOXJ1 gene expression (by RT-PCR), which was restored in a dose-dependent manner by NAC treatment (Mata et al., 2012). In another study, FOXJ1 mRNA levels were consistently low after RSV infection and were restored with roflumilast N-oxide (Mata et al., 2013). Thoracic irradiation reduced FOXJ1 mRNA levels in mouse lungs (Bernard et al., 2012). Irradiation causes excessive levels of free radicals and associated lipid peroxidation, damage to DNA, proteins, leading to wide-spread cellular damage (Azzam et al., 2012; Koc et al., 2003; Rodrigues-Moreira et al., 2017; Shirazi et al., 2013). The expression of cilia-related transcription factor genes, including FOXJ1, RFX2, and RFX3 was significantly down-regulated by CSE treatment. The expression of cilia motility and structural integrity genes, including DNAI1, DNAH5, DNAH9, DNAH10, DNAH11, and SPAG6 was also significantly down-regulated by CSE treatments (Brekman et al., 2014). Many of these genes (RFX2, RFX3, DNAI1, DNAH9, DNAH11, SPAG6) are transcriptionally regulated by FOXJ1 (Causal biological network database, 2019). The downregulation of FOXJ1-controlled genes infer reduced FOXJ1 transcription factor activity. Indeed, overexpression of FOXJ1 led to partial restoration of CSE treatment-induced downregulation of cilia-related genes (Brekman et al., 2014).

Schamberger et al. did not find any alterations in FOXJ1 mRNA levels or FOXJ1 target gene (DNAI1, DNALI1, SPAG6, TEKT1) transcription upon exposure of HBECs to 2.5% or 5% CSE for 28 days. However, in this study, cigarette smoke exposure reduced ciliated cell numbers (Schamberger et al., 2015). The evidences listed suggest several mechanisms on how oxidative stress could lead to decreased FOXJ1 levels, including EGFR-, MCIDAS- or IL-13-mediated mechanisms. Most of the studies, however, do not corroborate on how oxidative stress mechanistically leads to reduced FOXJ1 levels. Since there are several other factors (GMNC, NOTCH, ULK4 etc.) known to regulate FOXJ1 levels, further pathways might be involved in passing the oxidative stress signal to FOXJ1.

Unknown

High oxidative stress causes reduction in FOXJ1 levels measured at 24 h and for up to 15 days after exposure to an oxidative stress-causing agent. The data on cigarette smoke-reduced FOXJ1 levels are convincing. Indirect evidence such as antioxidants restoring CS-reduced FOXJ1 levels suggest that oxidative stress plays a major role in the CS-induced effects (Milara et al., 2012). However, given the complexity of the CS mixtures (Baumung et al., 2016), we cannot exclude that factors other than oxidative stress are involved in FOXJ1 level reduction. Other sources of oxidative stress such as RSV infection or GY radiation reduce FOXJ1 levels to similar degree as CS-caused FOXJ1 reduction. Based on the available evidence, we classify the quantitative understanding of this KER as moderate.

Normal HBECs were exposed to whole cigarette smoke from 3R4F research grade cigarettes using the Vitrocell® VC 10® Smoking Robot (35-mL puff volume, 2 s duration and 1 min between puffs or air as a control). Differentiated cells were exposed every 2 d for 5 d (3 exposures), and samples were collected 48 h after treatment. Differentiating cells were exposed 3 times per week to smoke from 1 cigarette, and samples were collected after 14, 21 and 27 days. FOXJ1 protein and mRNA levels decreased 2.5-fold in differentiated and 2-fold in differentiating normal HBECs. There was a significantly lower percentage of FOXJ1-positive cells following whole smoke exposure at day 27 (4.3 +/- 4.2% vs. 13.0 +/- 7.3%, air) (Valencia-Gattas et al., 2016). CSE was obtained from one Marlboro Red commercial cigarette bubbled in 12.5 mL of differentiation medium that was then filtered (0.2-µm pore filter). The absorbance was measured at 320 nm on a spectrophotometer, and the optical density of 1 was defined as 100% CSE. HBECs were differentiated at the air-liquid interface while being exposed to 0, 3, and 6% CSE between days 5 and 28. Treatment with 3% and 6% CSE reduced FOXJ1 mRNA levels to approx. 65% and 55% of control levels, respectively. Treatment of differentiating cultures with 3% CSE reduced FOXJ1 protein levels by 2-fold by day 28 (Brekman et al., 2014). The smoke of one 2R4F research cigarette was bubbled into a flask containing 25 mL of pre-warmed (37°C) differentiation medium using a respiratory pump model (Harvard Apparatus Rodent Respirator 680, Harvard Apparatus, Holliston, MA, USA) that generates three puffs/min; 35 mL per each puff of 2-s duration with a volume of 0.5 cm above the filter. The solution was then filtered (0.22 µm pore size) to remove particles and the tar phase. The resulting sterile solution was defined as 100% CSE and used within 30 min of preparation. Treatment of differentiated human bronchial epithelial cells with 10% CSE decreased FOXJ1 expression by about 40% at 24 h and 70% at 72 h exposure (Milara et al., 2012). 3R4F reference cigarettes were smoked in accordance with the ISO smoking protocol (35-mL puffs of 2 s each minute). Whole cigarette smoke, generated by a VC10 smoking robot, was released into a mixing device in 2.8-s exhaust and diluted with humidified clean air at 1.0 L/min dilution flow. Diluted smoke was introduced into the CULTEX RFS module and guided into the exposure chamber (5 mL/min) using a vacuum pump to expose 3D co-cultures of HBECs and fibroblasts. FOXJ1 mRNA levels were reduced to 60% and 40% of the control after exposure to CS from 1 or 4 cigarettes, respectively (Ishikawa and Ito, 2017). RSV infection elicits reactive oxygen intermediate-mediated effects manifested by changes in the expression of NRF2 and HMOX-1 genes (approx. 6- and 9-fold increase at day 15 post-RSV infection, respectively), H2O2 generation (7-fold increase in intracellular levels at day 15 post-infection) and severe reduction in total antioxidant capacity. These data together indicate the presence of oxidative stress following infection which leads to decreased FOXJ1 mRNA levels (ca. 25% of control at 15 days post-RSV infection (Mata et al., 2012) and ca. 45% of control at 10 days after RSV infection (Mata et al., 2013). FOXJ1 mRNA levels were reduced by 50% in murine lungs 14 days after thoracic irradiation at 15 Gy (Bernard et al., 2012).

Whole cigarette smoke exposure from one 3R4F research grade cigarette using the Vitrocell® VC 10® Smoking Robot (35-mL puff volume, 2-s duration and 1 min between puffs or air as a control) once a day on alternate days for 5 days decreased FOXJ1 mRNA levels by 2.5-fold in differentiated HBECs (Valencia-Gattas et al., 2016). Whole cigarette smoke exposure from one 3R4F research grade cigarette using the Vitrocell® VC 10® Smoking Robot (35-mL puff volume, 2-s duration and 1 min between puffs or air as a control) for 3 times per week for 4 weeks (27 days) decreased FOXJ1 mRNA levels by 2-fold in differentiating HBECs (Valencia-Gattas et al., 2016). Treatment of differentiating HBECs (between days 5 and 28 of the air-liquid interface) with 3% CSE reduced FOXJ1 protein levels by 2-fold by day 28. Treatment of differentiating HBECs with 6% CSE reduced FOXJ1 protein levels by approx. 55% by day 28 (Brekman et al., 2014). Treatment of differentiated human bronchial epithelial cells with 10% CSE decreased FOXJ1 expression by about 40% at 24 h and ca. 70% at 72 h (Milara et al., 2012).  Repeated exposure of 3D bronchial epithelial cultures to whole smoke of 4 cigarettes (every other day, treatment started on ALI culture day 7) resulted in a 2.5-fold decrease of FOXJ1 mRNA levels by day 21 (Ishikawa and Ito, 2017). At 15 days post-RSV infection, FOXJ1 mRNA levels were 4-fold reduced compared to untreated samples (Mata et al., 2012). In another study from the same group, FOXJ1 mRNA levels were reduced to 45% of the FOXJ1 levels in uninfected sample at 10 days post-RSV infection (Mata et al., 2013). At 7 and 14 days after 15 GY thoracic irradiation, FOXJ1 mRNA levels were reduced to approx. 70% and 50% of controls, respectively (Bernard et al., 2012).

Unknown

Not Specified Mixed

Not Specified

All life stages

High

Akaike, T., Ando, M., Oda, T., Doi, T., Ijiri, S., Araki, S., et al. (1990). Dependence on O2- generation by xanthine oxidase of pathogenesis of influenza virus infection in mice. J. Clin. Invest. 85, 739-745.  Azzam, E.I., Jay-Gerin, J.P., and Pain, D. (2012). Ionizing radiation-induced metabolic oxidative stress and prolonged cell injury. Cancer Lett. 327, 48-60.  Baumung, C., Rehm, J., Franke, H., and Lachenmeier, D.W. (2016). Comparative risk assessment of tobacco smoke constituents using the margin of exposure approach: the neglected contribution of nicotine. Sci. Rep. 6, 35577. Bernard, M.E., Kim, H., Rajagopalan, M.S., Stone, B., Salimi, U., Rwigema, J.C., et al. (2012). Repopulation of the irradiation damaged lung with bone marrow-derived cells. In Vivo 26, 9-18. Brekman, A., Walters, M.S., Tilley, A.E., and Crystal, R.G. (2014). FOXJ1 prevents cilia growth inhibition by cigarette smoke in human airway epithelium in vitro. Am. J. Respir. Cell Mol. Biol. 51, 688-700. Garcia-Arcos, I., Geraghty, P., Baumlin, N., Campos, M., Dabo, A.J., Jundi, B., et al. (2016). Chronic electronic cigarette exposure in mice induces features of COPD in a nicotine-dependent manner. Thorax 71, 1119-1129.  Ishikawa, S., and Ito, S. (2017). Repeated whole cigarette smoke exposure alters cell differentiation and augments secretion of inflammatory mediators in air-liquid interface three-dimensional co-culture model of human bronchial tissue. Toxicol. In Vitro 38, 170-178. Koc, M., Taysi, S., Buyukokuroglu, M.E., and Bakan, N. (2003). Melatonin protects rat liver against irradiation-induced oxidative injury. J. Radiat. Res. 44, 211-215.  Mata, M., Martinez, I., Melero, J.A., Tenor, H., and Cortijo, J. (2013). Roflumilast inhibits respiratory syncytial virus infection in human differentiated bronchial epithelial cells. PLoS One 8, e69670.  Mata, M., Sarrion, I., Armengot, M., Carda, C., Martinez, I., Melero, J.A., et al. (2012). Respiratory syncytial virus inhibits ciliagenesis in differentiated normal human bronchial epithelial cells: effectiveness of N-acetylcysteine. PloS One 7, e48037. Milara, J., Armengot, M., Bañuls, P., Tenor, H., Beume, R., Artigues, E., et al. (2012). Roflumilast N-oxide, a PDE4 inhibitor, improves cilia motility and ciliated human bronchial epithelial cells compromised by cigarette smoke in vitro. Br. J. Pharmacol. 166, 2243-2262.  Rodrigues-Moreira, S., Moreno, S.G., Ghinatti, G., Lewandowski, D., Hoffschir, F., Ferri, F., et al. (2017). Low-Dose Irradiation Promotes Persistent Oxidative Stress and Decreases Self-Renewal in Hematopoietic Stem Cells. Cell Rep. 20, 3199-3211.  Schamberger, A.C., Staab-Weijnitz, C.A., Mise-Racek, N., and Eickelberg, O. (2015). Cigarette smoke alters primary human bronchial epithelial cell differentiation at the air-liquid interface. Sci. Rep. 5, 8163. Shirazi, A., Mihandoost, E., Ghobadi, G., Mohseni, M., and Ghazi-Khansari, M. (2013). Evaluation of radio-protective effect of melatonin on whole body irradiation induced liver tissue damage. Cell J. 14, 292-297. Valencia-Gattas, M., Conner, G.E., and Fregien, N.L. (2016). Gefitinib, an EGFR Tyrosine Kinase inhibitor, Prevents Smoke-Mediated Ciliated Airway Epithelial Cell Loss and Promotes Their Recovery. PloS One 11, e0160216.  Zhou, F., and Roy, S. (2015). SnapShot: Motile Cilia. Cell 162, 224 (e221).  AOPWiki 2021-08-02T03:04:14

2021-09-28T07:28:31

<upstream-id>381aa3f3-9bed-41f2-afe5-10ce883fe336</upstream-id> <downstream-id>087499da-ddbb-4dc3-ac4b-be9d387fcd06</downstream-id> Forkhead box J1 (FOXJ1) is a master regulator of motile ciliogenesis which is necessary and also sufficient to program cells to grow functional motile cilia (Vij et al., 2012; Zhou and Roy, 2015). Studies in different model organisms have shown that the loss of FOXJ1 results in a loss of motile cilia (Brody et al., 2000; Chenet al., 1998; Stubbs et al., 2008; Vij et al., 2012).

Homozygous null mutation of Foxj1 results in complete absence of cilia in mouse respiratory epithelium (Chen et al., 1998; Brody et al., 2000). In a previous study, wild-type mice had approximately 20% heavily ciliated cells in the proximal pulmonary epithelium, while explanted Foxj1-/- mouse trachea had no ciliated cells (Gomperts et al., 2004). Loss of FOXJ1 orthologs FoxJ1–4 in flatworm Schmidtea mediterranea results in loss of ciliation of the ventral epithelium which closely resembles the human airway epithelium (Rompolas et al., 2009; Vij et al., 2012). Loss of Foxj1 activity in Xenopus and zebrafish—through antisense morpholino oligonucleotides—reduces cilia formation, while, conversely, ectopic Foxj1 overexpression results in formation of multiple motile cilia (Stubbs et al., 2008; Yu et al., 2008). There is a strong correlation between FOXJ1 and expression of the FOXJ1 ciliogenesis program genes in zebrafish, Xenopus and mouse cells (Abedalthagafi et al., 2016). Treatment with cigarette smoke extract downregulates FOXJ1 mRNA and protein expression, which is accompanied by a reduction in cilia length and number in human bronchial epithelial cells in vitro (Milara et al., 2012; Brekman et al., 2014). This can be prevented by overexpression of FOXJ1 (Brekman et al., 2014) or treatment with roflumilast N-oxide, which reduces intracellular free radical levels and increases FOXJ1 mRNA and protein expression (Milara et al., 2012).

The requirement of FOXJ1 for cells to grow functional motile cilia was demonstrated in 1998 in a mouse model study where targeted disruption of FOXJ1 resulted in the absence of motile cilia in the respiratory epithelium, oviduct, haploid sperm, and choroid plexus (Chen et al., 1998). Subsequently, many research groups consistently showed FOXJ1 requirement for cilia growth in various model organisms (Brody et al., 2000; Gomperts et al., 2007; Stubbs et al., 2008; Vij et al., 2012; Yu et al., 2008). In addition, overexpression of FOXJ1 in ectopic locations prompted cilia growth (Stubbs et al., 2008; Yu et al., 2008), and FOXJ1 overexpression could rescue cigarette smoke extract-caused cilia growth suppression in human airway epithelium (Brekman et al., 2014). The causal association of FOXJ1 to ciliogenesis gene expression program was computationally reinforced (Abedalthagafi et al., 2016). Taken together, the empirical support for this KER based on the research in the motile ciliogenesis field implies a high (strong) confidence for the biological plausibility of the linkage.

Two research groups independently showed that homozygous null mutation of Foxj1 causes complete absence of cilia in the respiratory airways of mice (Brody et al., 2000; Chen et al., 1998). Loss of Foxj1a activity in zebrafish (antisense morpholino oligonucleotides designed to block Foxj1a protein translation) compromises formation of motile cilia (Yu et al., 2008). In the same study, Foxj1a ectopic expression in the neural tube (through hyperactivation of the hedgehog pathway using dominant negative PKA) triggered motile cilia development (Yu et al., 2008). In Xenopus, FoxJ1 morpholino oligo treatments dose-dependently reduced cilia formation, and FoxJ1 overexpression in ectopic locations was sufficient to drive multiciliogenesis (Stubbs et al., 2008). In addition, authors also used a zebrafish model where FOXJ1 morpholino resulted in cilia number and length reduction (Stubbs et al., 2008). The closest ortholog of mammalian FOXJ1 in the flatworm Schmidtea mediterranea is FoxJ1-4 which is expressed in the motile ciliated cells of the ventral epithelium. Flatworm ventral epithelium closely resembles the mammalian airway epithelium (Rompolas et al., 2009; Vij et al., 2012). S. mediterranea deficient for foxJ1-4 (RNAi) lost the ciliation of the ventral epithelium (Vij et al., 2012). Abedalthagafi et al. defined the genes that comprise the FOXJ1 ciliogenesis program using public mRNA expression profiling data from Xenopus, zebrafish and mouse cells. A strong correlation of FOXJ1 expression and the expression of the FOXJ1 ciliogenesis program was confirmed (Abedalthagafi et al., 2016). Cigarette smoke extract (CSE) down-regulated FOXJ1 mRNA and protein levels and cilia length in differentiating human airway basal cells in air–liquid interface (ALI) cultures, while lentivirus-mediated FOXJ1 expression prevented CSE-elicited cilia growth suppression (Brekman et al., 2014). Wild-type mouse tracheas cultured at air-liquid interface demonstrated nearly 20% of the proximal pulmonary epithelial cells to be ciliated with abundant cilia on each cell. In explanted Foxj1-/- tracheas no ciliated epithelial cells were observed (Gomperts et al., 2004). Exposure of differentiated human bronchial epithelial cells to CSE decreased FOXJ1 expression alongside with reduction of the average number of cells with cilia motility. Roflumilast N-oxide (which reduced intracellular reactive oxygen species levels) prevented the CSE-elicited decrease in the expression levels of Foxj1 mRNA and protein. Concurrently, roflumilast N-oxide partly prevented the loss in cells with cilia motility (Milara et al., 2012). FOXJ1 overexpression increased the percentage of ciliated cells in mouse trachea organ culture 1.51-fold (Johnson et al., 2018).

Foxj1 overexpression failed to promote ciliogenesis in mouse polarized epithelial cell lines and primary cultured alveolar epithelial cells (You et al., 2004). Also, the overexpression of Foxj1 in wild-type airway epithelial cells did not enhance the total number of ciliated cells. However, delivery of Foxj1 to null cells resulted in cilia formation (You et al., 2004).

Regulatory factor X3 (RFX3) is a transcriptional co-activator of FOXJ1 (Didon et al., 2013) and is involved in motile cilia biogenesis (El Zein et al., 2009). Fluctuations in RFX3 levels can modulate the outcome that the upstream KE has on the downstream KE.

There is ample empirical evidence of FOXJ1 requirement for motile cilia formation, where complete removal of FOXJ1 results in cilia loss, and different levels of FOXJ1 downregulation result in proportional reduction in cilia number and length. Accordingly, FOXJ1 overexpression leads to higher cilia numbers. Based on these data of strong causality between upstream and downstream KEs, we judge our quantitative understanding to be high.

Complete removal of FOXJ1 by means of homologous recombination in mouse embryonic stem cells resulted in absence of cilia in mouse airways (as well as in other typically multiciliated tissues such as oviduct, haploid sperm, choroid plexus and epithelial cells of the brain but not in embryonic node) (Brody et al., 2000; Chen J. et al., 1998). Newly fertilized zebrafish eggs were injected with antisense morpholino oligonucleotides designed to block Foxj1a protein translation. Motile cilia numbers were severely reduced in Kupffer’s vesicle (KV), the ﬂoor plate and pronephric ducts 14 and 24 hpf (Yu et al., 2008). Downregulation of Xenopus FoxJ1 produced a dose-dependent defect in skin cilia formation. When 20 ng or 40 ng morpholino oligonucleotides were injected, cilia formed, but were reduced in number and shortened in length. After injection of 75 ng morpholino oligos, most cilia were lost. Cilia length decreased from ~11 microns to 4 microns (Stubbs et al., 2008). Morpholino oligo knockdown of zebrafish FoxJ1 caused a two-fold decrease in the number of KV cilia and a 3.5-fold decrease in the average length of KV cilia (Stubbs et al., 2008). RNAi against Schmidtea mediterranea foxJ1-4 substantially reduced the expression levels of foxJ1-4 which lead to almost complete loss of motile cilia (Vij et al., 2012). In the presence of CSE, in FOXJ1 overexpressing human airway epithelial cells the average cilia length was significantly higher (5.2 μm) than in lentivirus-control–infected cells (4.1 μm) (Brekman et al., 2014). CSE was obtained from one Marlboro Red commercial cigarette bubbled in 12.5 ml of differentiation medium that was then 0.2 mm pore filtered. The absorbance was measured at 320 nm on a spectrophotometer and the optical density of 1 was defined as 100%. Homozygous FOXJ1 mutant mice were obtained by mating foxj1+/- male and female animals. The explanted trachea of the foxj1-/- mice harbors no motile cilia in contrast to wild-type trachea (Gomperts et al., 2004). Exposure of differentiated human bronchial epithelial cells to 10% CSE decreased FOXJ1 expression by about 40% at 24 h and 70% at 72 h exposure. The smoke of one 2R4F research cigarette was bubbled into a flask containing 25 mL of pre-warmed (37°C) differentiation medium using a respiratory pump model (Harvard Apparatus Rodent Respirator 680, Harvard Apparatus, Holliston, MA, USA) that generates three puffs min−1; 35 mL per each puff of 2 s duration with a volume of 0.5 cm above the filter. The CS solution was filtered (0.22 µm pore size) to remove particles and the tar phase. The resulting sterile solution was defined as 100% CSE and used within 30 min of preparation. Exposure to CSE concentration- and time-dependently reduced the average number of cells with cilia motility, which was significant after 3 days of incubation with CSE at 2.5% (about 30% inhibition), and reached a maximum of about 75% inhibition versus control after 7 days of incubation with CSE at 10% (Milara et al., 2012). Roflumilast N-oxide at 2 nM or 1 µM concentration-dependently prevented the decrease in the expression levels of Foxj1 mRNA and protein following 3 days of exposure of differentiated bronchial epithelial cells to CSE at 10%. Concurrently, roflumilast N-oxide partly prevented the loss in cells with cilia motility (Milara et al., 2012). Electroporation using negative control (GFP-only) plasmid resulted in 45±1.4% (mean±s.e.m.) GFP+ ciliated cells in mouse trachea organ culture. FOXJ1 significantly increased the percentage of GFP+ ciliated cells to 68±3.6% (1.51-fold) (Johnson et al., 2018).

14- or 24-hr treatment with antisense morpholino oligonucleotides designed to block Foxj1a protein translation results in absence of motile cilia in zebrafish (Yu et al., 2008). Xenopus embryos were injected with FOXJ1 morpholino oligos at two-cell stage (1.5 h of embryo life) and the embryos were analyzed at stage 26 (1 day, 5 h and 30 min of embryo life) for cilia phenotype (Stubbs et al., 2008). Schmidtea mediterranea worms received three feedings of foxJ1-4 RNAi (2 days in between feeds) and were analyzed for cilia phenotype 14 days after the last feed. RNAi against Schmidtea mediterranea foxJ1-4 substantially reduced the expression levels of foxJ1-4 which lead to almost complete loss of motile cilia (Vij et al., 2012). Exposure of differentiated human bronchial epithelial cells to CSE concentration- and time-dependently reduced the average number of cells with cilia motility, which was significant after 3 days of incubation with CSE at 2.5% (about 30% inhibition), and reached a maximum of about 75% inhibition versus control after 7 days of incubation with CSE at 10% (Milara et al., 2012).

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<ul> <li>Abedalthagafi, M.S., Wu, M.P., Merrill, P.H., Du, Z., Woo, T., Sheu, S.H., et al. (2016). Decreased FOXJ1 expression and its ciliogenesis programme in aggressive ependymoma and choroid plexus tumours. J. Pathol. 238(4), 584-597.</li> <li>Brekman, A., Walters, M.S., Tilley, A.E., and Crystal, R.G. (2014). FOXJ1 prevents cilia growth inhibition by cigarette smoke in human airway epithelium in vitro. Am. J. Respir. Cell Mol. Biol. 51(5), 688-700.</li> <li>Brody, S.L., Yan, X.H., Wuerffel, M.K., Song, S.K., and Shapiro, S.D. (2000). Ciliogenesis and left-right axis defects in forkhead factor HFH-4-null mice. Am. J. Respir. Cell Mol. Biol. 23(1), 45-51.</li> <li>Chen, J., Knowles, H.J., Hebert, J.L., and Hackett, B.P. (1998). Mutation of the mouse hepatocyte nuclear factor/forkhead homologue 4 gene results in an absence of cilia and random left-right asymmetry. J. Clin. Investig. 102(6), 1077-1082. </li> <li>Didon, L., Zwick, R.K., Chao, I.W., Walters, M.S., Wang, R., Hackett, N.R., et al. (2013). RFX3 Modulation of FOXJ1 regulation of cilia genes in the human airway epithelium. Respir. Res. 14(1), 70-70. </li> <li>El Zein, L., Ait-Lounis, A., Morle, L., Thomas, J., Chhin, B., Spassky, N., et al. (2009). RFX3 governs growth and beating efficiency of motile cilia in mouse and controls the expression of genes involved in human ciliopathies. J Cell Sci 122(Pt 17), 3180-3189. </li> <li>Gomperts, B.N., Gong-Cooper, X., and Hackett, B.P. (2004). Foxj1 regulates basal body anchoring to the cytoskeleton of ciliated pulmonary epithelial cells. J. Cell Sci. 117(Pt 8), 1329-1337. </li> <li>Gomperts, B.N., Kim, L.J., Flaherty, S.A., and Hackett, B.P. (2007). IL-13 Regulates Cilia Loss and foxj1 Expression in Human Airway Epithelium. Am. J. Respir. Cell Mol. Biol. 37(3), 339-346. </li> <li>Johnson, J.A., Watson, J.K., Nikolic, M.Z., and Rawlins, E.L. (2018). Fank1 and Jazf1 promote multiciliated cell differentiation in the mouse airway epithelium. Biol Open 7(4). bio033944. </li> <li>Milara, J., Armengot, M., Bañuls, P., Tenor, H., Beume, R., Artigues, E., et al. (2012). Roflumilast N-oxide, a PDE4 inhibitor, improves cilia motility and ciliated human bronchial epithelial cells compromised by cigarette smoke in vitro. Br. J. Pharmacol. 166(8), 2243-2262. </li> <li>Rompolas, P., Patel-King, R.S., and King, S.M. (2009). Schmidtea mediterranea: a model system for analysis of motile cilia. Methods Cell Biol. 93, 81-98. </li> <li>Stubbs, J.L., Oishi, I., Izpisua Belmonte, J.C., and Kintner, C. (2008). The forkhead protein Foxj1 specifies node-like cilia in Xenopus and zebrafish embryos. Nat. Genet. 40(12), 1454-1460. </li> <li>Vij, S., Rink, J.C., Ho, H.K., Babu, D., Eitel, M., Narasimhan, V., et al. (2012). Evolutionarily ancient association of the FoxJ1 transcription factor with the motile ciliogenic program. PLoS Genet. 8(11), e1003019. d</li> <li>You, Y., Huang, T., Richer, E.J., Schmidt, J.-E.H., Zabner, J., Borok, Z., et al. (2004). Role of f-box factor foxj1 in differentiation of ciliated airway epithelial cells. American Journal of Physiology-Lung Cellular and Molecular Physiology 286(4), L650-L657. </li> <li>Yu, X., Ng, C.P., Habacher, H., and Roy, S. (2008). Foxj1 transcription factors are master regulators of the motile ciliogenic program. Nat. Genet. 40(12), 1445-1453.</li> <li>Zhou, F., and Roy, S. (2015). SnapShot: Motile Cilia. Cell 162(1), 224-224 e221. </li> </ul> AOPWiki 2021-07-23T06:40:11

2021-08-02T10:08:02

<upstream-id>087499da-ddbb-4dc3-ac4b-be9d387fcd06</upstream-id> <downstream-id>93a4ab2a-7008-4c39-afc2-d20fd61c6998</downstream-id> The cilia in the respiratory epithelium beat in a coordinated fashion at a frequency of approximately 10 to 16 Hz, propelling mucus upwards (Joki et al., 1998; Smith et al., 2012). Many factors, including cilia length, number, structure, orientation as well as mucus viscosity, temperature, pH, chemicals, airway surface liquid height, exposure to bacterial and viral pathogens have been shown to affect ciliary function (Clary-Meinesz et al., 1998; Ho et al., 2001b; Jing et al., 2017; Joki et al., 1998; Kanthakumar et al., 1996; Mall, 2008; Smith et al., 2012; Snyder et al., 2017). Alteration from normal physiological conditions and from healthy cilia number/length/structure typically reduces the cilia beat frequency (CBF) (Clary-Meinesz et al., 1998; Jayathilake et al., 2012).

In Chlamydomonas, ciliary motion is directly related to the length of the cilia (Bottier et al., 2018). Similar observations have been made in zebrafish, where modulation of cilia length and number by FOR20 (centrosomal protein 20) deletion/knockdown directly impairs ciliary motility (Xie et al., 2019). There is also a positive correlation between cilia number and CBF in sinusitis patients (Joki et al., 1998), while cilia number, length and orientation correlate positively with mucociliary transport rate in patients with recurrent or longstanding respiratory infections (Toskala et al., 1995; Joki et al., 1998). Comparisons of strips of normal and disrupted ciliated epithelium have shown that CBF is decreased in the latter (Thomas et al., 2009). Mathematical models and simulations have shown that periciliary liquid and mucus velocity are directly affected by cilia number and length (Lee et al., 2011; Jayathilake et al., 2012; Jayathilake et al., 2015).

Many research groups have shown positive correlations between cilia number/length and ciliary function in studies dating as early as 1995. In some cases, the authors measured mucociliary clearance as the endpoint, thus the evidence does not describe causality between cilia number/length and CBF. However, the commonly held assumption is that reduced ciliary function, i.e., reduced CBF, leads to decreased mucociliary clearance. Based on the evidence, we judge the biological plausibility of this KER as moderate.

Cilia beating frequency analysis in sinusitis patient samples showed a positive correlation between cilia number and CBF (Joki et al., 1998). Cilia study from patients with recurrent or longstanding respiratory infections demonstrated a correlation between mucociliary transport rate and cilia number, length, and orientation (Toskala et al., 1995). According to a numerical model formulated by Jayathilake et al., the average periciliary liquid layer (PCL) velocity decreases when cilia are shortened (Jayathilake et al., 2015; Jayathilake et al., 2012). A study of Chlamydomonas cilia motion concluded that cilia shorter than 4 µm show variable or no periodicity of beating. For cilia longer than 4 µm, the ratio of frequency estimated from body motion to frequency of cilia motion is very close to 1.0, as expected. In short (2 µm, 4 µm) cilia, the ratio was significantly different from 1; apparently, even if the short cilium beats periodically, the frequency of beating is not reliably transmitted to the body motion (Bottier et al., 2018). In a simulation study by Lee et al., an increase in cilia number increased mucus velocity (Lee et al., 2011). In zebrafish, deletion or knockdown of FOR20 caused reduced number and length of cilia. Ciliary motility is impaired in animals with reduced FOR20 (Xie et al., 2019). Cilia beating frequency measurements on normal ciliated or disrupted epithelial strips of tissue revealed lower CBF for tissues with cilia disruptions (Thomas et al., 2009).

Although the majority of studies discuss the effect of shorter cilia on ciliary function, there are also reports on longer than normal cilia impairing cilia function (Jayathilake et al., 2015; Li et al., 2014b). A range of factors other than cilia length and number can influence cilia beat frequency. Often a combination of two or more factors affect ciliary function making it difficult to discern the impact of an individual factor on CBF (Toskala et al., 1995; Xie et al., 2019).

Unknown

Cilia beating frequency correlates with cilia size and numbers such that higher number of motile cilia with a healthy length show efficient CBF and a reduction of cilia number and/or length results in a proportionate reduction of CBF. In some studies, mucociliary clearance was measured as an indicator of CBF. Based on evidence presented here, we judge the quantitative understanding to be moderate.

A study in nasal mucosa samples from sinusitis patients demonstrated that reduced cilia numbers account for decreased CBF. In normally ciliated samples the CBF was 11.2±3.7 Hz, in samples with some ciliated cells CBF was 8.9±6.3 Hz, and in samples with no detectable cilia CBF was 2.1±3.8 Hz (Joki S. et al., 1998). Patients with recurrent or longstanding respiratory infections were divided into 3 groups based on mucociliary transport rates (MTR). The group with slowest MTR had the biggest number of non-ciliated columnar cells in the nasal mucosa, and the highest number of short and disoriented cilia. Loss of ciliated cells was seen in 50% of specimens with good MTR (7 to 10.8 mm per minute), in 71% with moderate MTR (3 to 6.9 mm per minute) and in 86% with poor MTR (0 to 2.9 mm per minute). The percentages of short cilia were 1% in the first group, 6% in the second, and 10% in the third. In this study, ciliary disorientation also contributed to the low MTR (Toskala et al., 1995). Jayathilake et al. developed a two-dimensional numerical model for computing how the length of cilia is affecting fluid flow. In the model, the cilia length was reduced by 5%, 15%, 25%, and 35% from 6 µm while other parameters were kept unchanged. The average periciliary layer (PCL) velocity in stream-wise direction decreases when cilia are shortened. When the cilia are shortened by 10%, the average stream-wise PCL velocity is reduced by about 11%, while the average span-wise PCL velocity is reduced by about 62% (Jayathilake et al., 2012). In a similar study by the same group, the length of cilia was defined as 50%, 60%, 70%, 80%, 90%, 100%, 110%, 120%, 130% and 140% of the length of the healthy cilia (i.e., 6 µm). The mucus velocity reaches its maximum value when the ciliary length is around the length of the healthy cilia (or standard length of 6 µm) (Jayathilake et al., 2015). Cilia shorter than 4 µm have anomalies in the ciliary beating periodicity. Cilia shorter than 2 µm were never found to beat periodically. Cilia between 2 and 4 µm in length exhibited more variable periodicity. Cilia with lengths between 4 and 12 µm beat periodically with conserved frequency (Bottier et al., 2018). Mucus velocity increased in concordance with cilia number increase. Mean mucus velocity increased from 26.82 µm/s to 49.53 µm/s when cilia number grew from 10 to 24 (Lee et al., 2011). FOR20 deletion or knockdown in zebrafish reduced the number and length of cilia. Cilia in FOR20 morphants had impaired ciliary motility. When cilia length was reduced from an average 5.5 µm to an average 3 µm due to FOR20 depletion, about 80% cilia displayed consistently paralyzed or arrhythmically beating pattern (Xie et al., 2019). Nasal brush biopsy samples from pediatric patients with primary ciliary dyskinesia were analyzed. The median CBF was 13.4 Hz for normal ciliated epithelia, 11.4 Hz for the ciliated edge with minor projections and 8.7 Hz for the ciliated edge with major projections (Thomas et al., 2009).

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High Not Specified

Bottier, M., Thomas, K., Dutcher, S., and Bayly, P. (2018). (Preprint) How does cilium length affect beating? bioRxiv, 474346.  Clary-Meinesz, C., Mouroux, J., Cosson, J., Huitorel, P., and Blaive, B. (1998). Influence of external pH on ciliary beat frequency in human bronchi and bronchioles. Eur. Respir. J. 11(2), 330-333. Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The Effect of Aging on Nasal Mucociliary Clearance, Beat Frequency, and Ultrastructure of Respiratory Cilia. Am. J. Respir. Crit. Care Med. 163(4), 983-988. Jayathilake, P.G., Le, D.V., Tan, Z., Lee, H.P., and Khoo, B.C. (2015). A numerical study of muco-ciliary transport under the condition of diseased cilia. Comput. Methods Biomech. Biomed. Engin. 18(9), 944-951.  Jayathilake, P.G., Tan, Z., Le, D.V., Lee, H.P., and Khoo, B.C. (2012). Three-dimensional numerical simulations of human pulmonary cilia in the periciliary liquid layer by the immersed boundary method. Comput. Fluids 67, 130-137. Jing, J.C., Chen, J.J., Chou, L., Wong, B.J.F., and Chen, Z. (2017). Visualization and Detection of Ciliary Beating Pattern and Frequency in the Upper Airway using Phase Resolved Doppler Optical Coherence Tomography. Sci. Rep. 7(1), 8522-8522.  Joki, S., Toskala, E., Saano, V., and Nuutinen, J. (1998). Correlation between ciliary beat frequency and the structure of ciliated epithelia in pathologic human nasal mucosa. Laryngoscope 108(3), 426-430.  Kanthakumar, K., Taylor, G., Cundell, D., Dowling, R., Johnson, M., Cole, P., et al. (1996). The effect of bacterial toxins on levels of intracellular adenosine nucleotides and human ciliary beat frequency. Pulm. Pharmacol. 9(4), 223-230. Lee, W., Jayathilake, P., Tan, Z., Le, D., Lee, H., and Khoo, B. (2011). Muco-ciliary transport: effect of mucus viscosity, cilia beat frequency and cilia density. Comput. Fluids 49(1), 214-221. Li, Y.Y., Li, C.W., Chao, S.S., Yu, F.G., Yu, X.M., Liu, J., et al. (2014). Impairment of cilia architecture and ciliogenesis in hyperplastic nasal epithelium from nasal polyps. J. Allergy Clin. Immunol. 134, 1282-1292.  Mall, M.A. (2008). Role of cilia, mucus, and airway surface liquid in mucociliary dysfunction: lessons from mouse models. J. Aerosol Med. Pulm. Drug Deliv. 21, 13-24. Smith, C.M., Djakow, J., Free, R.C., Djakow, P., Lonnen, R., Williams, G., et al. (2012). ciliaFA: a research tool for automated, high-throughput measurement of ciliary beat frequency using freely available software. Cilia 1, 14.  Snyder, R.J., Hussain, S., Tucker, C.J., Randell, S.H., and Garantziotis, S. (2017). Impaired Ciliogenesis in differentiating human bronchial epithelia exposed to non-Cytotoxic doses of multi-walled carbon Nanotube. Part. Fibre Toxicol. 14, 1-14.  Thomas, B., Rutman, A., and O'Callaghan, C. (2009). Disrupted ciliated epithelium shows slower ciliary beat frequency and increased dyskinesia. Eur. Respir. J. 34, 401-404.  Toskala, E., Nuutinen, J., Rautiainen, M., and Torkkeli, T. (1995). The correlation of mucociliary transport and scanning electron microscopy of nasal mucosa. Acta Otolaryngol. 115, 61-65.  Xie, S., Jin, J., Xu, Z., Huang, Y., Zhang, W., Zhao, L., et al. (2019). Centrosomal protein FOR20 is essential for cilia-dependent development in zebrafish embryos. FASEB J. 33, 3613-3622.  AOPWiki 2021-07-23T06:40:39

2021-09-28T07:41:47

<upstream-id>93a4ab2a-7008-4c39-afc2-d20fd61c6998</upstream-id> <downstream-id>1e732e79-9f2e-4a4e-a83a-39a51ee3ccc8</downstream-id> Synchronized ciliary action transports mucus from the distal lung to the mouth, where it is swallowed or expectorated (Munkholm and Mortensen, 2014). In addition to ASL and mucus properties, the speed of ciliary movement, and hence the effectiveness of mucociliary clearance (MCC), is dependent on ciliary amplitude and beat frequency (Rubin, 2002). CBF itself is influenced by several factors, including changes in the physical and chemical properties of the ASL (especially the periciliary fluid), structural modulation in the cilia, concentration of cyclic nucleotides cAMP and cGMP, and intracellular calcium (Ca2+). Aside from genetic defects leading to ciliopathies, there is ample evidence for prolonged exposure to noxious agents, such as cigarette smoke, nitrogen oxide and sulfur dioxide, playing a major role in decreasing CBF and hampering efficient MCC.

A decrease in CBF resulting from sulfur dioxide exposure reduced mucociliary clearance in dogs (Yeates et al., 1997) and mucociliary activity in guinea pig tracheas (Knorst et al., 1994). In rats, formaldehyde inhalation exposure resulted in lower numbers of ciliated cells, while ciliary activity and mucus flow rates were decreased in a dose and time-dependent manner (Morgan et al., 1986). In humans, CBF positively correlates with nasal mucociliary clearance time (Ho et al., 2001), and bronchiectasis patients have lower nasal CBF and slower mucociliary transport (MCT) (Rutland and Cole, 1981). Administration of nebulized CBF inhibitors and enhancers quantifiably decreased or increased mucociliary clearance, respectively (Boek et al., 1999; Boek et al., 2002). Increased CBF and MCT was also noted in human sinonasal epithelial cell cultures treated with Myrtol®, an essential oil distillate (Lai et al., 2014) and in sheep tracheas and human airway epithelial cultures subjected to temperature changes (Kilgour et al., 2004; Sears et al., 2015). Exposures of frog palate epithelia to formaldehyde and PM10 reduced MCC and mucociliary transport, but only formaldehyde-treated epithelia showed decreases in CBF (Morgan et al., 1984; Macchione et al., 1999; Fló-Neyret et al., 2001). Ex vivo treatment of sheep trachea with acetylcholine and epinephrine increased CBF, but only acetylcholine increased surface liquid velocity, while both parameters were decreased upon incubation with platelet-activating factor (Seybold et al., 1990).

Ciliary function and mucus transport are invariably linked to effective mucus transport along the mucociliary escalator (Bustamante-Marin and Ostrowski, 2017; Mall, 2008). Therefore, this KER is biologically plausible.

Studies in animal models of ciliopathies and in individuals with genetic disorders causing cilia defects demonstrate that absent or asynchronous cilia beating results in defective mucus clearance from the lungs, consequently leading to respiratory infections that may be chronic recurrent in nature and ultimately lead to declining lung function (Knowles et al., 2013; Munkholm and Mortensen, 2014; Tilley et al., 2015). Similarly, indirect effects of airway inflammation, caused for example by respiratory infections or allergies, are known to be responsible for changes in cilia beating and hence mucus clearance (Almeida-Reis et al., 2010; Hisamatsu and Nakajima, 2000; Maurer et al., 1982). Finally, airway epithelial injury following exposure to inhalation toxicants can also damage cilia and inhibit cilia function and thereby impair MCC (Iravani and Van As, 1972; Wanner et al., 1996). A decrease in CBF resulting from sulfur dioxide exposure reduced mucociliary clearance in dogs (Yeates et al., 1997) and mucociliary activity in guinea pig tracheas (Knorst et al., 1994). In rats, formaldehyde inhalation exposure resulted in lower numbers of ciliated cells, while ciliary activity and mucus flow rates were decreased in a dose and time-dependent manner (Morgan et al., 1986). In humans, CBF positively correlates with nasal mucociliary clearance time (Ho et al., 2001), and bronchiectasis patients have lower nasal CBF and slower mucociliary transport (MCT) (Rutland and Cole, 1981). Administration of nebulized CBF inhibitors and enhancers quantifiably decreased or increased mucociliary clearance, respectively (Boek et al., 1999; Boek et al., 2002). Increased CBF and MCT was also noted in human sinonasal epithelial cell cultures treated with Myrtol®, an essential oil distillate (Lai et al., 2014) and in sheep tracheas and human airway epithelial cultures subjected to temperature changes (Kilgour et al., 2004; Sears et al., 2015). Exposures of frog palate epithelia to formaldehyde and PM10 reduced MCC and mucociliary transport, but only formaldehyde-treated epithelia showed decreases in CBF (Morgan et al., 1984; Macchione et al., 1999; Fló-Neyret et al., 2001). The available evidence does not interrogate the direct relationship between CBF and MCC, but rather evaluates both outcomes in parallel. However, because of the intrinsic linkage of cilia function and MCC, we find the empirical evidence in support of this KER to be moderate. Ex vivo treatment of sheep trachea with acetylcholine and epinephrine increased CBF, but only acetylcholine increased surface liquid velocity, while both parameters were decreased upon incubation with platelet-activating factor (Seybold et al., 1990).

Although ciliary function is considered a primary determinant for effective MCC (Duchateau et al., 1985; Gizurarson, 2015), there is evidence that suggests that MCC can be impeded by other factors that do not affect CBF. For example, nasal CBF in cigarette smokers regularly exhaling through the nose was not significantly different from that of nonsmokers, although they exhibited significantly longer nasomuciliary clearance times compared to nonsmokers. Possible explanations offered for this discrepancy were a potential loss of cilia in the nasal epithelium or increased mucus viscoelasticity (Stanley et al., 1986). Similarly, formaldehyde exposure of rats resulted in decreased cilia numbers and slower mucus flow rates (Morgan KT et al., 1986). On the other hand, there are a number of pharmacological compounds that improve mucociliary clearance through reductions in mucus viscosity, but have no effect on CBF (Jiao and Zhang, 2019), or through increases in CBF, but have no effect on mucociliary clearance (Phillips et al., 1990).

Physiological factors such as age, sex, posture, sleep, and exercise were shown to affect MCC, although study findings are not always concordant (Houtmeyers et al., 1999). MCC and CBF, for example, were observed to decrease with age in several species in numerous studies (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Yager et al., 1978), but evidence by (Agius et al., 1998) suggests that age does not have a major effect on CBF.

There are several studies providing insights into the negative effect of inhalation exposures on CBF and MCC, that are in line with the current thinking on how these two KEs connect. Additionally, pharmacological studies demonstrated that stimulation of CBF typically results in stimulation of MCC. However, since most studies usually evaluated the KEs in parallel, and even though some results support both dose response and temporal sequence of the KEs, none of the available data affirms causal linkage between CBF and MCC. Our understanding of the evidence is therefore moderate.

CBF decreased sequentially with increasing SO2 doses in dogs. CBF decreased from 6.3 ± 0.2 (SE) Hz at baseline to 5.7 ± 0.2 Hz at 5.5 ppm SO2. Five ppm SO2 delivered to both the trachea and tracheobronchial airways for 20 min also caused a marked decrease in mean bronchial mucociliary clearance from 53.7 ± 5.7% to 32.8 ± 7.7% after 90 min (Yeates et al., 1997). The effects of 30-min exposure to SO2 on mucociliary activity (MCA) and ciliary beat frequency (CBF) were studied in 31 guinea pig tracheas. A 63% reduction in mean MCA and statistically insignificant changes in CBF were recorded at concentrations of 2.5 ppm SO2. Higher SO2 concentrations caused further impairment of MCA as well as a dose-dependent decrease in CBF: At 5 ppm SO2, CBF decreased by 45%, at 12.5 ppm by 72%. The maximum decrease in MCA (81%) was observed with 7.5 ppm SO2; the highest SO2 concentration did not decrease MCA further. The decrease in MCA was associated with an impairment of CBF only at SO2 concentrations ≥5.0 ppm (Knorst et al., 1994b). Administration of a nebulized CBF inhibitor (0.9% NaCl) to 15 healthy volunteers significantly decreased mucociliary transport (MCT) from 7.9±1.5 mm/min (SEM) to 4.5±1.6 mm/min. Salbutamol, a CBF enhancer, significantly increased MCT from 8.0±1.4 to 12.5±1.1 mm/min (Boek et al., 2002; Boek et al., 1999). Cooling human airway epithelial cultures grown at the air-liquid interface from 37°C to 25°C over the course of approx. 20 min decreased CBF from 12 to 6 Hz and mucociliary transport (MCT) from 140 to 90 µm/s. Extending the range of temperature tested, CBF was found to increase by 0.49±0.06 Hz for every temperature increase by 1°C, and this was mirrored by an increase in MCT. MCT increased on average between 5 and 11 µm/s for every Hz increase in CBF. This study also showed that CBF decreased with increasing mucin concentration, dropping from 12.4 Hz at 2% bovine submaxillary mucin (BSM) to 10.1 Hz at 8% BSM, concurrent with a ca. 70% reduction in MCT. In addition, treatment with 10 µM basolateral forskolin reproducibly increased CBF by 19.3±2.1% and MCT by 24.4±3.1% over baseline (Sears et al., 2015). In sheep trachea CBF and mucus transport velocity (MTV) were 9.8±2.7 beats/s and 5.7±2.6 mm/min, respectively, at baseline. Temperature reductions from 37°C to 34°C caused a progressive decline in CBF (ca. –20% at 2 h and –90% at 4 h) and MTV (ca. –50% at 2 h and –90% at 4 h), which was further exacerbated by additional temperature decreases (30°C; CBF: ca. –75% at 2 h; MTV: –80% at 2 h) (Kilgour et al., 2004). Frog palate preparations were incubated with 1.25, 2.5 and 5.0 ppm formaldehyde. At formaldehyde doses of 2.5 and 5 ppm, CBF decreased by ca. 25% compared to baseline within 30 min and by 35-50% within 60 min (Fló-Neyret et al., 2001). Incubation of frog palates with PM10 from Sao Paolo, Brazil, for up to 120 min did not affect CBF but decreased MCT at concentrations ≥1000 pg/m3 (Macchione et al., 1999) In freshly excised sheep tracheas, a 60-min incubation with 10 µM platelet-activating factor caused a 6% decrease in CBF and a dose-dependent decrease in surface liquid velocity, reaching a maximum of 63% (Seybold et al., 1990). In patients with bronchiectasis, nasal CBF was 12.8±1.3 Hz and nasal clearance time was 31.8± 18.4 min. In comparison, in healthy controls, nasal CBF was 14.0±1.3 Hz and nasal clearance time was 17.6± 8.3 min (Rutland and Cole, 1981). Following basolateral treatment of human sinonasal epithelial cell cultures grown at the air-liquid interface with  Myrtol®, a phytopharmaceutical mixture of distillates of rectified essential oils of eucalyptus, sweet orange, myrtle, and lemon as the active ingredients, increased CBF in a dose-dependent manner, with a maximum stimulation with 0.1% of 48±7% after 30 min. The same concentration caused a 46±16% increase in MCT at 40 min (Lai et al., 2014). In New Zealand white rabbits exposed to 3 ppm NO2 for 24 h, the average CBF decreased from 764 beats/min to 692 beats/min, and the transport velocity decreased from 5.23 mm/min to 3.03 mm/min (Kakinoki, 1998).

A 20-minute exposure of dogs to SO2 caused a decrease in mean bronchial MCC after 90 min (Yeates et al., 1997). Frog palate epithelia were incubated with 1.25, 2.5 and 5.0 ppm formaldehyde. At formaldehyde doses of 2.5 and 5 ppm, CBF decreased by ca. 25% compared to baseline within 30 min and by 35-50% within 60 min (Fló-Neyret et al., 2001). Incubation of freshly excised sheep tracheas with 10 µM platelet-activating factor caused a maximal decrease in CBF of 6% after 60 min and decrease in surface liquid velocity of ca. 30% at 20 min, ca. 50% at 40 min and 63% after 60 min (Seybold et al., 1990). Following basolateral treatment of human sinonasal epithelial cell cultures grown at the air-liquid interface with different concentrations of Myrtol®, CBF increased rapidly within the first 30 min and then declined thereafter. The maximum response for MCT was seen after 40 min (Lai et al., 2014).

Unknown

High Mixed

High

All life stages

High Not Specified Not Specified Not Specified Not Specified Not Specified

Evidences for this KER are derived from studies carried out in dog, gunea pig, rat, frog, sheep, rabbit model systems as well as in human epithelial cell cultures. MCC and CBF were observed to decrease with age in several species (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Yager et al., 1978), but evidence by (Agius et al., 1998) suggests that age does not have a major effect on CBF.

Agius, A.M., Smallman, L.A., and Pahor, A.L. (1998). Age, smoking and nasal ciliary beat frequency. Clin. Otolaryngol. Allied Sci. 23, 227-230.  Almeida-Reis, R., Toledo, A.C., Reis, F.G., Marques, R.H., Prado, C.M., Dolhnikoff, M., et al. (2010). Repeated stress reduces mucociliary clearance in animals with chronic allergic airway inflammation. Respir. Physiol. Neurobiol. 173, 79-85. Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., DeVasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589.  Boek, W.M., Graamans, K., Natzijl, H., van Rijk, P.P., and Huizing, E.H. (2002). Nasal Mucociliary Transport: New Evidence for a Key Role of Ciliary Beat Frequency. Laryngoscope 112, 570-573.  Boek, W.M., Keleş, N., Graamans, K., and Huizing, E.H. (1999). Physiologic and hypertonic saline solutions impair ciliary activity in vitro. Laryngoscope 109, 396-399. Bustamante-Marin, X.M. and Ostrowski, L.E. (2017). Cilia and Mucociliary Clearance. Cold Spring Harb. Persp. Biol. 9, a028241. Duchateau, G.S., Merkus, F.W., Zuidema, J., and Graamans, K. (1985). Correlation between nasal ciliary beat frequency and mucus transport rate in volunteers. The Laryngoscope 95, 854-859. Fló-Neyret, C., Lorenzi-Filho, G., Macchione, M., Garcia, M.L.B., and Saldiva, P.H.N. (2001). Effects of formaldehyde on the frog's mucociliary epithelium as a surrogate to evaluate air pollution effects on the respiratory epithelium. Braz. J. Med. Biol. Res. 34, 639-643. Gizurarson, S. (2015). The effect of cilia and the mucociliary clearance on successful drug delivery. Biol. Pharmaceut. Bull. b14-00398. Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B., and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867. Hisamatsu, K.-i., and Nakajima, M. (2000). Pranlukast protects leukotriene C4- and D4-induced epithelial cell impairment of the nasal mucosa in vitro. Life Sci. 67, 2767-2773.  Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The Effect of Aging on Nasal Mucociliary Clearance, Beat Frequency, and Ultrastructure of Respiratory Cilia. Am. J. Respir. Crit. Care Med. 163, 983-988.  Houtmeyers, E., Gosselink, R., Gayan-Ramirez, G., and Decramer, M. (1999). Regulation of mucociliary clearance in health and disease. Eur. Respir. J. 13, 1177-1188. Iravani, J., and Van As, A. (1972). Mucus transport in the tracheobronchial tree of normal and bronchitic rats. J. Pathol. 106, 81-93. Jiao, J., and Zhang, L. (2019). Influence of Intranasal Drugs on Human Nasal Mucociliary Clearance and Ciliary Beat Frequency. Allergy Asthma Immunol. Res. 11, 306-319.  Joki, S., and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea‐pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169. Kakinoki, Y.O., Ayaki Tanaka, Yushi Washio, Koji Yamada, Yoshiaki Nakai, Kazuhiro Morimoto, Yasushi (1998). Nitrogen dioxide compromises defence functions of the airway epithelium. Acta Otolaryngol. 118, 221-226. Kilgour, E., Rankin, N., Ryan, S., and Pack, R. (2004). Mucociliary function deteriorates in the clinical range of inspired air temperature and humidity. Intensive Care Med. 30, 1491-1494. Knorst, M.M., Kienast, K., Riechelmann, H., Müller-Quernheim, J., and Ferlinz, R. (1994). Effect of sulfur dioxide on mucociliary activity and ciliary beat frequency in guinea pig trachea. Int. Arch. Occup. Environ. Health 65, 325-328. Knowles, M.R., Daniels, L.A., Davis, S.D., Zariwala, M.A., and Leigh, M.W. (2013). Primary ciliary dyskinesia. Recent advances in diagnostics, genetics, and characterization of clinical disease. Am. J. Respir. Crit. Care Med. 188, 913-922. Lai, Y., Dilidaer, D., Chen, B., Xu, G., Shi, J., Lee, R.J., et al. (2014). In vitro studies of a distillate of rectified essential oils on sinonasal components of mucociliary clearance. Am. J. Rhinol. Allergy 28, 244-248. Macchione, M., Oliveira, A.P., Gallafrio, C.T., Muchão, F.P., Obara, M.T., Guimarães, E.T., et al. (1999). Acute effects of inhalable particles on the frog palate mucociliary epithelium. Environ. Health Perspect. 107), 829-833. Mall, M.A. (2008). Role of cilia, mucus, and airway surface liquid in mucociliary dysfunction: lessons from mouse models. J. Aerosol Med. Pulm. Drug Delivery 21, 13-24. Maurer, D., Sielczak, M., Oliver Jr, W., Abraham, W., and Wanner, A. (1982). Role of ciliary motility in acute allergic mucociliary dysfunction. J. Appl. Physiol. 52, 1018-1023. Morgan, K., Patterson, D., and Gross, E. (1986). Responses of the nasal mucociliary apparatus of F-344 rats to formaldehyde gas. Toxicol. Appl. Pharmacol. 82, 1-13. Munkholm, M., and Mortensen, J. (2014). Mucociliary clearance: pathophysiological aspects. Clin. Physiol. Funct. Imaging 34, 171-177. Paul, P., Johnson, P., Ramaswamy, P., Ramadoss, S., Geetha, B., and Subhashini, A.S. (2013). The Effect of Ageing on Nasal Mucociliary Clearance in Women: A Pilot Study. ISRN Pulmonology 2013, 5.  Phillips, P.P., McCaffrey, T.V., and Kern, E.B. (1990). The in vivo and in vitro effect of phenylephrine (Neo Synephrine) on nasal ciliary beat frequency and mucociliary transport. Otolaryngology Head Neck Surg. 103, 558-565. Rubin, B.K. (2007). Mucus structure and properties in cystic fibrosis. Paediatr. Respir. Rev. 8, 4-7.  Rutland, J., and Cole, P.J. (1981). Nasal mucociliary clearance and ciliary beat frequency in cystic fibrosis compared with sinusitis and bronchiectasis. Thorax 36, 654-658. Sears, P.R., Yin, W.-N., and Ostrowski, L.E. (2015). Continuous mucociliary transport by primary human airway epithelial cells in vitro. Am. J. Physiol. Lung Cell. Mol. Physiol. 309, L99-L108.  Seybold, Z.V., Mariassy, A.T., Stroh, D., Kim, C.S., Gazeroglu, H., and Wanner, A. (1990). Mucociliary interaction in vitro: effects of physiological and inflammatory stimuli. J. Appl. Physiol. 68, 1421-1426.  Stanley, P., Wilson, R., Greenstone, M., MacWilliam, L., and Cole, P. (1986). Effect of cigarette smoking on nasal mucociliary clearance and ciliary beat frequency. Thorax 41, 519-523. Tilley, A.E., Walters, M.S., Shaykhiev, R., and Crystal, R.G. (2015). Cilia dysfunction in lung disease. Ann. Rev. Physiol. 77, 379-406.  Wanner, A., Salathe, M., and O'Riordan, T.G. (1996). Mucociliary clearance in the airways. Am. J. Respir. Crit. Care Med. 154, 1868-1902.  Yager, J., Chen, T.-M., and Dulfano, M.J. (1978). Measurement of frequency of ciliary beats of human respiratory epithelium. Chest 73, 627-633. Yeates, D.B., Katwala, S.P., Daugird, J., Daza, A.V., and Wong, L.B. (1997). Excitatory and inhibitory neural regulation of tracheal ciliary beat frequency (CBF) activated by ammonia vapour and SO2. Ann. Occup. Hyg. 41, 736-744. AOPWiki 2021-07-19T10:24:02

2023-03-24T08:17:45

<upstream-id>1e732e79-9f2e-4a4e-a83a-39a51ee3ccc8</upstream-id> <downstream-id>94c02ede-6ad2-4283-8e73-99ca863b484a</downstream-id> It is very well known that patients suffering from motile ciliopathies, such as primary ciliary dyskinesia, have impaired or absent MCC and lower lung function (reduced FEV1 and FVC) compared to their healthy counterparts (Halbeisen et al., 2018; Marthin et al., 2010; Wallmeier et al., 2020). In cystic fibrosis patients, decreased MCC (due to reduced airway hydration and changes in mucus chemical and viscoelastic properties) causes mucus build-up leading to mucus plugging in the airways and consequently to decreased lung function over time (Kerem et al., 2014; Mossberg et al., 1978; Regnis et al., 1994; Robinson and Bye, 2002; Szczesniak et al., 2017; Wanner et al., 1996). Mucus plugging due to decreased MCC is also considered a major cause of airway obstruction and airflow limitation in COPD patients (Dunican et al., 2021; Okajima et al., 2020) and asthmatics (Kuyper et al., 2003; Maxwell, 1985).

Changes in MCC rate are typically paralleled by effects on lung function in several studies where both endpoints have been assessed. In patients with primary ciliary dyskinesia, absence of cilia motion prevents normal MCC and consequently, lung function is reduced (Denizoglu Kulli et al., 2020). In cystic fibrosis patients, the ASL is depleted resulting in impaired MCC (Boucher, 2004). Although the known CFTR genotypes can result in a variety of phenotypes (Derichs, 2013), clinical data indicate that some specific gene defects, such as the p.Phe508del variant, are more frequently associated with decreased lung function indices (e.g. FEV1 % predicted, FVC % predicted, FEF25-75) (Kerem et al., 1990; Johansen et al., 1991; Schaedel et al., 2002). Both cigarette smoking and occupational exposure to biomass fumes led to slower MCC and reduced FEV1 % predicted and FEV1/FVC (Ferreira et al., 2018). Nasomucociliary clearance was slower in COPD smokers compared to former smokers with COPD or to nonsmokers (Ito et al., 2015). Allergen challenge in asthma patients resulted in both reduced MCC and FEV1, which could be reversed by inhalation of hypertonic saline solution (Alexis et al., 2017). In cystic fibrosis patients, treatment with mucolytic agents (Laube et al., 1996; McCoy et al., 1996; Quan et al., 2001; Elkins et al., 2006; Amin et al., 2011; Donaldson et al., 2018) or a CFTR potentiator (Rowe et al., 2014) improved both MCC and lung function (FEV1, FVC and FEF25-75).

Lung function is known to decrease with age, and several studies showed that mucus transport rates also decrease in older compared to younger individuals (Goodman et al., 1978; Uzeloto et al., 2021). Impaired MCC is also seen in chronic smokers, even prior to a clinically significant drop in lung function and the detection of small airway disease (Clunes et al., 2012a; Goodman et al., 1978; Lourenço et al., 1971; Uzeloto et al., 2021; Vastag et al., 1986), and in patients with obstructive lung disease and hence, poor lung function (Cruz et al., 1974; Vastag et al., 1986). Adult asthmatics also displayed decreased mucus transport rates/velocities in addition to decreased lung function (Ahmed et al., 1981; Bateman et al., 1983; Foster et al., 1982; Mezey et al., 1978). In patients with primary ciliary dyskinesia, absence of cilia motion prevents normal MCC and consequently, lung function is reduced (Denizoglu Kulli et al., 2020). In cystic fibrosis patients, the ASL is depleted resulting in impaired MCC (Boucher, 2004a). Although the known CFTR genotypes can result in a variety of phenotypes (Derichs, 2013), clinical data indicate that some specific gene defects, such as the p.Phe508del variant, are more frequently associated with decreased lung function indices (e.g. FEV1 % predicted, FVC % predicted, FEF25-75) (Johansen et al., 1991; Kerem et al., 1990; Schaedel et al., 2002). Unsurprisingly, results from studies with pharmacological agents aimed at restoring CFTR function do not only indicate enhanced MCC but also support improvements in lung function (Bennett et al., 2018; Donaldson et al., 2018; Rowe S. M. et al., 2014a). While the available data link these two KEs, causal evidence is not always available, and some inference is present. Therefore, we judge the biological plausibility of this KER as moderate.

Occupational exposure to biomass combustion products resulted in slower MCC and reduced FEV1 % predicted and FEV1/FVC (Ferreira et al., 2018). Compared to healthy controls, current smokers without airway obstruction and current smokers with COPD exhibited longer saccharin transit times, indicative of impaired MCC, and lower FEV1 % predicted and FEV1/FVC (Uzeloto et al., 2021). Similarly, nasomucociliary clearance was slower in COPD smokers compared to former smokers with COPD or to nonsmokers (Ito et al., 2015). Additionally, mucus plug density—assessed by CT imaging—and mucoid (rather than watery) consistency were inversely related to FEF25–75% and associated with increased RV/TLV (Kesimer et al., 2018). Asthma patients responded to allergen challenge with a reduction in both MCC and FEV1 (Bennett et al., 2011; Mezey et al., 1978), which could be rescued by inhalation with hypertonic saline solution (Alexis et al., 2017). Multiple studies interrogating the effect of mucolytic agents such as hypertonic saline solution or recombinant DNase on mucus transport rates or mucus clearance in patients with cystic fibrosis report improvements in both, mucus transport velocities or rates and lung function indices, including FEV1, FVC and FEF25-75 (Amin et al., 2011; Donaldson et al., 2018; Elkins et al., 2006; Laube et al., 1996; McCoy et al., 1996; Quan et al., 2001). Both MCC and lung function (FEV1, FVC and FEF25-75) improved in cystic fibrosis patients treated with ivacaftor, a CFTR potentiator that increases the channel open probability (Rowe et al., 2014b). Some studies with mucolytics such as N-acetylcysteine, bromhexine, theophylline/ambroxol or sorebrol demonstrated improved MCC was connected with small improvements in lung function (FEV1, FVC and FEV1/FVC) in patients with chronic bronchitis (Aylward et al., 1980; Castigiioni and Gramolini, 1986; Thomson et al., 1974; Würtemberger et al., 1988).

Genetic defects leading to motile ciliopathies or defects in CFTR function are linked to impaired MCC. However, because of the genetic variety, not every defect, for example in the CFTR gene, also expresses an overt pulmonary phenotype. Other factors, such as low-level chronic inflammation may drive lung pathology by pathways independent of MCC. This might also explain the absence of differences in MCC between healthy smokers and smokers with COPD (Fleming et al., 2019). Not all studies looking to elucidate the effect of mucolytics on MCC report an improvement of lung function, even though mucus transport rates or tracheobronchial clearance significantly improve. These studies include, for example, some on the effects of hypertonic saline solution, NAC, ambroxol and 2-mercapto-ethane sulphonate (Clarke et al., 1979; Ericsson et al., 1987; Millar et al., 1985; Robinson et al., 1997; Würtemberger et al., 1988). This could be, at least in part, related to the fact that a sudden drop in lung function served as an indicator of patient distress in these studies, and interventions were halted when they occurred to ensure patient safety (Robinson et al., 1996). Another reason could be related to the mechanisms underlying mucus solubilization that may be completely independent of lung function. MCC is only one means by which mucus can be cleared from the lungs. Another one is cough clearance, and it is highly dependent on the properties of the ASL, in particular the ASL height (Knowles and Boucher, 2002).

Invariably, if mucus viscosity increases (independent of whether that results from increased mucus production (hypersecretion), depletion of the ASL or another cause) and MCC decreases, another mechanism comes into action to clear excess mucus: cough clearance. Cough constitutes a “backup” host defense by which acutely or chronically accumulated mucus is expelled through forceful, high-velocity airflow (Button et al., 2018; King, 2006). Our current understanding of the mechanical principles and biology of cough suggest that failure of cough clearance may also be a contributor to decreased lung function.

The available data, though not causally linking decreases in MCC with decreased lung function, provide a good insight into the importance of the physiological role of MCC in maintaining normal lung function. In at least some studies, impairment of MCC correlated with the drop in FEV1 or FEF25-75. Although clinically valuable benefits can be seen in studies with pharmacological agents such as mucolytics and CFTR modifying drugs, they do not cover a wide range of dose responses nor are they supportive of the KER causality. Therefore, we judge our quantitative understanding as moderate.

Sixteen Brazilian sugarcane workers aged 25±4 years, with a BMI of 24±3 kg/m2, with exhaled CO of 2.1±1.5 ppm, were examined during the non-harvest season and during the sugarcane burning harvest season. There was a non-significant decrease in saccharin transit time (from 8±1 min to 3±1 min) and a significant decrease  FEV1/FVC ratio (from 88.62±5.68 to 84.90±6.47) and %FEV1 (from 92.19±13.24 to 90.44±12.76) during harvest compared with the non-harvest season (Ferreira et al., 2018). 12 (6M/6F) mild allergic, non-smoking asthmatics ages 20–39 with skin sensitivity to house dust mites (HDM) and normal baseline lung function (FEV1 %pred > 80, FEV1/FVC ratio >0.70) inhaled sequential doses of inhaled HDM extract (D. farinae, Greer®, Lenoir, NC) delivered as 5 inhalations from a Devilbiss 646 nebulizer (mass median aerodynamic diameter of 5 um, GSD = 2.0). Five of the 12 patients responded to the allergen challenge with >10% reductions in FEV1 % predicted and reduction in whole lung MCC as evidenced by increased retention rates (mean Central TB Ave120Ret increased from 0.69 to 0.79 for baseline vs. allergen challenge respectively). This reduction in MCC significantly correlated with the post challenge 24 hour FEV1 (Bennett et al., 2011). Treatment of patients with chronic bronchitis with bromhexine (3 x 16 mg/day) for 14 days resulted in mean changes in FEV1, FVC and FEV1/FVC of + 0.047 L + 0.033 L and +0.6%, respectively, with MCC at 6 h being 6.8% greater after treatment compared to baseline (Thomson et al., 1974). Treatment of patients with chronic bronchitis with ambroxol alone (2 x 30 mg/day) or with theophyllin (2 x 400 mg/day) and ambroxol (2 x 30 mg/day) for 7 days MCC/h improved from 18.3 ± 11.1% to 23.3 ± 13% and 29.6 ± 15.7%, respectively whereas lung function remained nearly unchanged with FEV1 predicted of 86.0 ± 9.78 at baseline vs 83.7 ± 9.27 (ambroxol only) and 83.1 ± 11.07 (combination) (Würtemberger et al., 1988). Treatment of chronic bronchitics with N-acetylcysteine (4 mg/day by metered dose inhaler) for 16 weeks significantly improved sputum viscosity (-0.53 vs -0.67; differences between medians to placebo: 0-14 (-0.77 0.64)) and minimally improved FVC (3.0±0.21 vs 2.9± 0.18 L/s) and PEF (356.7 ±29.64 L/min vs 354.6±25.07) but not FEV1 (1.9±0.18 vs 2.0± 0.13 L/s) (Dueholm et al., 1992). Treatment of asthmatics with salmeterol improved tracheobronchial clearance rates (AUC: 333±24%h vs 347±30%h in placebo) as well as FEV1 (76 ± 8), FVC (100 ± 5) and PEF % predicted (100 ± 7) compared to placebo (73 ± 8; 95 ± 5; 94 ± 7) (Hasani et al., 2003). Treatment of mild-to-moderate bronchitics with 42 µg salmeterol slightly enhanced whole lung clearance in 2 hr (not significant; C10–2= 25±11% vs 22±10% in placebo), significantly increased mean  peripheral lung clearance (C10–2= 22±9% vs 17±10% in placebo) and significantly increased FEV1 %pred and FEF25–75 at 2 h compared to baseline (93±18%predicted, 2.45 ± 1.08 L/s vs 88±19%predicted, 2.27 ± 0.98 L/s in placebo) (Bennett et al., 2006). Sputum induction by inhalation of hypertonic saline solution (5%) in asthmatics at 6 hr following challenge with LPS significantly improved FEV % predicted by approx. 20% and was accompanied by a ca. 6-fold increase in whole lung clearance (from 0.1 %/min to 0.6%/min) (Alexis et al., 2017). 133 cystic fibrosis patients (age (mean [SD]) was 21.1 (11.4) years and 46.4% were female. All participants had one copy of the G551D mutation, and 72.2% were compound heterozygous with F508del on the other allele.) completed a 6-month course of ivacaftor. Lung function improved from baseline FEV1% predicted of 82.6 (25.6) to 90.1 (25.0) (mean change, 6.7; 95% CI, 4.9–8.5). In a subgroup of 22 patients, particle clearance from the whole right lung was markedly increased. Average clearance through 60 minutes at 1 month post-treatment was more than twice the baseline value, reflecting substantially improved MCC (Rowe et al., 2014b). Inhalation of hypertonic saline solution (7%, 4 mL twice daily for 48 weeks) by cystic fibrosis patients improved FVC (by 82 mL; 95 percent confidence interval, 12 to 153) and FEV1 (by 68 mL; 95 percent confidence interval, 3 to 132) values, but not FEF25–75 (Elkins et al., 2006). In cystic fibrosis patients that inhaled hypertonic saline solution without amiloride twice a day over a period of 14 days one-hour mucus clearance rates improved from baseline (9.3±1.6%) to 14.0±2.0% and increased FEV1 by 6.2%. FVC and FEF25-75 also improved by 1.8% and 13.1%, respectively (Donaldson et al., 2006). Dornase alfa (recombinant human DNase) is currently used as a mucolytic to treat pulmonary disease in cystic fibrosis. It reduces mucus viscosity in the lungs, promoting improved clearance of secretions (Yang and Montgomery, 2021). In children with cystic fibrosis (mean: 8.4 yrs of age with FEV1 ≥95% predicted) treated with dornase alfa for 96 weeks, FEV1 % predicted improved by 3.2 ± 1.2, FVC % predicted improved by 0.7 ± 1.0, and FEF25-75 % predicted improved by 7.9 ± 2.3 compared to placebo (Quan et al., 2001). In young patients with cystic fibrosis (6-18 yrs of age with FEV1 ≥80% predicted) treated with dornase alfa for 96 weeks, FEF25-75 % predicted improved by 6.1±10.34 compared to placebo (Amin et al., 2011). In 10 adult cystic fibrosis patients receiving 2.5 mg rhDNase twice a day for 6 days, FEV1 and FVC increased by an average of 9.4 ± 3.5% and 12.7 ± 2.6%, respectively, as compared with a decrease of 1.8 ± 1.7% and an increase of 0.4±1.1% in the placebo group, respectively, although there were no significant changes in MCC (Laube et al., 1996). In 320 cystic fibrosis patients (7 to 57 yrs of age), dornase alfa treatment at 2.5 mg/day for 12 weeks (McCoy et al., 1996). Saccharin transit times (a marker of nasal MCC) were higher in healthy current smokers and COPD smokers than in healthy controls (10.87 [7.29–17] min and 16.47 [8.25–20.15] min, respectively, vs 8.52 [5.54–13.91] min). These groups also differed in their lung function indices: FEV1 % predicted was 101.4 ± 12.37 in healthy controls, 96.41 ± 12.3 in healthy current smokers, and 67.96 ± 24.02 in COPD smokers. FVC % predicted was 103.1 ± 13.45 in healthy controls, 97.51 ± 12.88 in healthy current smokers, and 90.33 ± 29.27 in COPD smokers. FEV1/FVC % predicted was 82.15 [78.5–85] in healthy controls, 82.20 [79.2–84.1] in healthy current smokers, and 61.1 [55.3–67.2] in COPD smokers (Uzeloto et al., 2021). Saccharin transit time of smokers with COPD (16.5 [11–28] min, median [interquartile range 25–75%]) was slightly longer than that of current smokers (15.9 [10 –27] min), and both were longer compared with exsmokers with COPD (10.2 [6 –12] min) and nonsmokers (8 [6 –16] min). Lung function parameters for the groups were as follows: nonsmokers, FEV1/FVC 0.84 ± 0.09, FEV1 % predicted  103.2 ± 11.5, FVC % predicted 102.2 ± 13.3; current smokers, FEV1/FVC 0.76 ± 0.05, FEV1 % predicted  90.7 ± 7.4, FVC % predicted 96.3 ± 13.9; former smokers with COPD, FEV1/FVC 0.49 ± 0.08, FEV1 % predicted  46.8 ± 12.6, FVC % predicted 76.8 ± 18.5; current smokers with COPD, FEV1/FVC 0.66 ± 0.16, FEV1 % predicted  48.7 ± 16.8, FVC % predicted 71.7 ± 13.0 (Ito et al., 2015).

Six asymptomatic patients with bronchial asthma and a history of allergic pollenosis and episodic bronchospasm consistent with ragweed hypersensitivity wer challenged by inhalation of an aqueous, short ragweed antigen extract (Greer Laboratories, Lenoir, N.C.), diluted with a phosphate-buffered saline solution. Mean tracheal mucus velocity (TMV) decreased to 72% of baseline immediately after challenge when specific airway conductance (SGaw), and FEV1 showed a maximal decrease, with a further decrease to 47% of baseline after 1 h, when SGaw and FEV1 had returned to baseline values (Mezey et al., 1978). Treatment of chronic bronchitics with N-acetylcysteine (3 x 200 mg/day) for 4 weeks significantly decreased sputum thickness, increased sputum pourability from 650% glycerol time (at baseline) to 320% glycerol time on day 21 and PEFR on days 28 (+5%), 35 (+6%) and 42 (+7%) and FEV1 on days 21 (+2%), 28(+3%), 35 (+4%) and 42 (+5%) compared to baseline (ca. 33% predicted and 28% predicted, respectively) (Aylward et al., 1980). Treatment of mild-to-moderate bronchitics with 42 µg salmeterol slightly enhanced whole lung clearance in 2 hr (not significant; C10–2= 25±11% vs 22±10% in placebo), significantly increased mean  peripheral lung clearance (C10–2= 22±9% vs 17±10% in placebo) and significantly increased FEV1 %pred and FEF25–75 at 2 h compared to baseline (93±18%predicted, 2.45 ± 1.08 L/s vs 88±19%predicted, 2.27 ± 0.98 L/s in placebo) , and significantly increased FEV1 %pred and FEF25–75 at both 1 (92±19%predicted, 2.44 ± 1.14 L/s) and 2 h (93±18%predicted, 2.45 ± 1.08 L/s) compared to baseline (pre-dose; 90 ± 20%predicted, 2.16 ± 0.92 L/s) (Bennett et al., 2006). In cystic fibrosis patients on a 6-month ivacaftor regimen, FEV1% improvement was detectable as soon as the 1-month follow-up visit (mean change, 6.7; 95% CI, 5.2–8.3) (Rowe et al., 2014b). MCC remained at elevated level at the month 3 visit (Donaldson et al., 2018). One-hour mucus-clearance rates in cystic fibrosis patients receiving hypertonic saline with placebo were significantly faster than in the group receiving hypertonic saline with amiloride (14.0±2.0 vs. 7.0±1.5 %), and the durability of response following the inhalation of hypertonic saline with placebo was ≥8 hours (Donaldson et al., 2006).

Unknown

High Mixed

High

All life stages

High

The evidences for this KER come from and therefore apply to humans.

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Fibros. 16, 318-326.  Thomson, M., Pavia, D., Gregg, I., and Stark, J. (1974). Bromhexine and mucociliary clearance in chronic bronchitis. Brit. J. Diseases Chest 68, 21-27. Uzeloto, J.S., Ramos, D., Silva, B.S.d.A., Lima, M.B.P.d., Silva, R.N., Camillo, C.A., et al. (2021). Mucociliary Clearance of Different Respiratory Conditions: A Clinical Study. Int. Arch. Otorhinolaryngol. 25, e35-e40.  Vastag, E., Matthys, H., Zsamboki, G., Köhler, D., and Daikeler, G. (1986). Mucociliary clearance in smokers. Eur. J. Respir. Dis. 68, 107-113. Wallmeier, J., Nielsen, K.G., Kuehni, C.E., Lucas, J.S., Leigh, M.W., Zariwala, M.A., et al. (2020). Motile ciliopathies. Nat. Rev. Dis. Prim. 6, 1-29. Würtemberger, G., Michaelis, K., and Matthys, H. (1988). [Additive action of theophylline and ambroxol on bronchial clearance?]. Prax. Klin. Pneumol. 42 Suppl 1, 300-303. AOPWiki 2021-07-19T10:24:31

2023-03-24T08:27:51

Oxidative Stress Leading to Decreased Lung Function via Decreased FOXJ1

ox stress-mediated FOXJ1/cilia/CBF/MCC impairment

Karsta Luettich

Karsta Luettich, Philip Morris Products S.A., Philip Morris International R&D, Neuchatel, Switzerland Hasmik Yepiskoposyan, Philip Morris Products S.A., Philip Morris International R&D, Neuchatel, Switzerland Monita Sharma, PETA Science Consortium International e.V., Stuttgart, Germany Frazer Lowe, Broughton Nicotine Services, Earby, Lancashire, United Kingdom Damien Breheny, British American Tobacco (Investments) Ltd., Group Research and Development, Southampton, United Kingdom

BY-SA

2.0

This AOP evaluates one of the major processes known to be involved in regulating efficient mucociliary clearance (MCC). MCC is a key aspect of the innate immune defense against airborne pathogens and inhaled chemicals and is governed by the concerted action of its functional components, the cilia and the airway surface liquid (ASL), which is composed of mucus and periciliary layers (Bustamante-Marin and Ostrowski, 2017). In response to various irritants and pathogens mucus is secreted by goblet cells, and cilia sweep mucus upward by coordinated beating motions thus clearing the airways from these substances. The ciliated airway epithelial cells are typically covered by hundreds of motile cilia. Cilia formation is initiated and coordinated by a distinct gene expression program, led by the transcription factor forkhead box J1 (FOXJ1) (Brody et al., 2000; Zhou and Roy, 2015). FOXJ1 appears to be the major factor in multiciliogenesis, whereby its activity is necessary and also sufficient for programming cells to assemble functional motile cilia (Vij et al., 2012). A decrease in the levels or absence of FOXJ1 protein in cells of the respiratory tract therefore inhibits ciliogenesis, preventing physiological mucus clearance and decreasing MCC. MCC dysfunction is linked to airway diseases such as chronic obstructive pulmonary disease (COPD) or asthma, both of which are characterized by decreased lung function and bear a significant risk of increased morbidity and mortality. With a surface area of ~100 m2 and ventilated by 10,000 to 20,000 liters of air per day (National Research Council, 1988; Frohlich et al., 2016), the lungs are a major barrier that protect the body from a host of external factors that enter the respiratory system and may cause lung pathologies. Mucociliary clearance (MCC) is a key aspect of the innate immune defense against airborne pathogens and inhaled particles and is governed by the concerted action of its functional components, the cilia and the airway surface liquid (ASL), which comprises mucus and the periciliary layer (Bustamante-Marin and Ostrowski, 2017). In healthy subjects, ≥10 mL airway secretions are continuously produced and transported daily by the mucociliary escalator. Disturbances in any of the processes regulating ASL volume, mucus production, mucus viscoelastic properties, or ciliary function can cause MCC dysfunction and are linked to airway diseases such as chronic obstructive pulmonary disease (COPD) or asthma, both of which bear a significant risk of increased morbidity and mortality. The mechanism by which exposure to inhaled toxicants might lead to mucus hypersecretion and thereby impact pulmonary function has already been mapped in AOP148 on decreased lung function. However, whether an exposure-related decline in lung function is solely related to excessive production of mucus is debatable, particularly in light of the close relationship between mucus, ciliary function, and efficient MCC. To date, no single event has been attributed to MCC impairment, and it is likely that events described in this AOP as well as in AOPs 148, 411 and 424 have to culminate to lead to decreased lung function.

Established regulatory guideline studies for inhalation toxicity focus on evident clinical signs of systemic toxicity, including death, or organ-specific toxicity following acute and (sub)chronic exposure respectively. In toxicological and safety pharmacological studies with airborne test items targeting the airways or the lungs as a whole, lung function is a relevant endpoint for the characterization of potential adverse events (OECD, 2018a; Hoymann, 2012). Hence, the AO “decreased lung function” is relevant for regulatory decision-making in the context of (sub)chronic exposure (OECD, 2018b; OECD, 2018c). Regulatory relevance of the AO “decreased lung function” is evident when looking at the increased risk of diseases in humans following inhalation exposure, and because of its links to other comorbidities and mortality. To aid diagnosis and monitoring of fibrosis, current recommendations include both the recording of potential environmental and occupational exposures as well as an assessment of lung function (Baumgartner et al., 2000). The latter typically confirms decreased lung function as demonstrated by a loss of lung volume. As the disease progresses, dyspnea and lung function worsen, and the prognosis is directly linked to the decline in FVC (Meltzer and Noble, 2008). Chronic exposure to cigarette smoke and other combustion-derived particles results in the development of COPD. COPD is diagnosed on the basis of spirometry results as laid out in the ATS/ERS Task Force documents on the standardization of lung function tests and their interpretation (Pellegrino et al., 2005; Culver et al., 2017, Graham et al., 2019). Rapid rates of decline in the lung function parameter FEV1 are linked to higher risk of exacerbations, increased hospitalization and early death (Wise et al., 2006; Celli, 2010). Reduced FEV1 also poses a risk for serious cardiovascular events and mortality associated with cardiovascular disease (Sin et al., 2005; Lee et al., 2015).

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The experimental evidence to support the biological plausibility of the KERs from MIE to AO is moderate to strong overall for the AOP presented here, while there is a moderate concordance of dose-response relationships. In terms of essentiality, we have rated all of the KEs as either moderate or high. AOPs such as this one can play a central role in risk assessment strategies for a wide variety of regulatory purposes by providing mechanistic support to an integrated approach to testing and assessment (IATA; (Clippinger et al., 2018)). IATAs are flexible frameworks that can be adapted to best address the regulatory question or purpose at hand. More specifically, this AOP can be applied to the risk assessment of inhaled toxicants, by enabling the development of testing strategies through the assembly of existing information and the generation of new data where they are currently lacking. Targeted approaches to fill data gaps can be developed using new approach methodologies (NAMs) informed by this AOP. All KE proposed in this AOP occur and are measurable in several species, including frogs, mice, rats, guinea pigs, ferrets, sheep, and humans. The majority of the supporting empirical evidence derives from studies in rodent and human systems, and experimental findings in animals appear to be highly translatable to humans. Data regarding the applicability of KE to all life-stages from birth to adulthood are available for the MIE (Oxidative Stress), KE2 (FOXJ1 Protein, Decreased), KE3 (Motile Cilia Number/Length, Decreased), KE4 (Cilia Beat Frequency, Decreased), KE5 (Mucociliary Clearance, Decreased), and AO (Decreased Lung Function), and indicate that they apply to all life stages. It is also worth noting here that age-dependent decreases in CBF, MCC, and lung function have been demonstrated in several species (e.g., guinea pigs, mice, and humans) and reflect normal physiological aging processes (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Sharma and Goodwin, 2006). Gender-specific data relevant to the AOP are not as widely available as species-specific data, and to our knowledge, the role of gender has not been systematically evaluated for all KE described here. Considering the essentiality of FOXJ1 for the ciliogenesis program and the impact of ciliary beating on MCC, we consider this AOP applicable to both genders.

The definition of essentiality implies that the modulation of upstream KEs impacts the downstream KEs in an expected fashion. If blocked or failing to occur, the KEs in the current AOP will not necessarily stop the progression to subsequent KEs. Due to the complex biology of motile cilia formation and function, ASL homeostasis, mucus properties and MCC, the KEs and AO may be triggered because of alternative pathways or biological redundancies. However, when exacerbated, the KEs promote the occurrence of downstream events eventually leading to the AO. The causal pathway starting from the exposure to oxidants and leading to decreased lung function involves parallel routes with KEs, each of which is sufficient to cause the downstream KE to occur. Different mechanisms, such as oxidant-induced decreases in ASL height via CFTR function decline (AOP424) or oxidant-induced decreases in cilia number and length as a result of decreased FOXJ1 levels, lead to decreased CBF and decreased MCC. Each of these pathways contributes to the AO, but their relative contributions are difficult to evaluate. Based on the evidence we judge the MIE (Oxidative Stress), KE2 (FOXJ1 Protein, Decreased), KE3 (Motile Cilia Number/Length, Decreased), KE4 (Cilia Beat Frequency, Decreased), and KE5 (Mucociliary Clearance, Decreased) highly essential.

We judge the overall biological plausibility of this AOP as strong. The KER Decreased FOXJ1 protein leading to decreased motile cilia length/number is supported by multiple studies across different species with ample empirical evidence reflecting both dose-response and time concordance. Other KER, such as Oxidative stress leading to decreased FOXJ1 lack this expanse of empirical evidence, or the evidence does not fully support the causality between the KE (Reduced cilia number/length leading to decreased CBF, Decreased CBF leading to decreased MCC) even though the relationship is logical and plausible.

Overall, our quantitative understanding of the AOP network is moderate. There is robust evidence that provides an insight into several KER presented here, and the dose response and temporal relationship between the two KE in question are well described and quantified for different stressors across different test systems (Decreased FOXJ1 protein leading to decreased motile cilia length/number; Decreased motile cilia length/number leading to decreased cilia beating frequency; Decreased cilia beat frequency leading to decreased MCC). In some instances, we are less confident in our quantitative understanding. For example, dose response data as well as data supportive of the KE causality are limited for the KER Decreased MCC leading to decreased lung function.

Given the individual and public health burden of the consequences of lung function impairment, gaining a greater understanding of the underlying mechanisms is extremely important in the risk assessment of respiratory toxicants. An integrated assessment of substances with the potential to be inhaled, either intentionally or unintentionally, could incorporate inhalation exposure and dosimetry modelling to inform an in vitro approach with appropriate exposure techniques and cell systems to assess KEs in this AOP (EPA’s Office of Chemical Safety and Pollution Prevention, 2019). Standardization and robustness testing of assays against explicit performance criteria using suitable reference materials can greatly increase the level of confidence in their use for KE assessment (Petersen et al., 2021). Much of the empirical evidence that supports the KERs in the qualitative AOP described here was obtained from in vitro studies using well-established methodologies for biological endpoint assessment. Being chemical agnostic, this AOP can be applied to a variety of substances that share the AO. For example, impaired MCC and decreased lung function have a long-known relationship with smoking, but little is known about the consequences of long-term use of alternative inhaled nicotine delivery products such as electronic cigarettes and heated tobacco products. This AOP can form the basis of an assessment strategy to evaluate the effects of exposure to aerosol from these products based on the KEs identified here.

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Antunes, M.B., and Cohen, N.A. (2007). Mucociliary clearance–a critical upper airway host defense mechanism and methods of assessment. Curr. Opin. Allergy Clin. Immunol. 7, 5-10. Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., DeVasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589. Bustamante-Marin, X.M., and Ostrowski, L.E. (2017a). Cilia and Mucociliary Clearance. Cold Spring Harb. Persp. Biol. 9, a028241. EPA’s Office of Chemical Safety and Pollution Prevention (2019). "FIFRA Scientific Advisory Panel Meeting Minutes and Final Report No. 2019-01 Peer Review on Evaluation of a Proposed Approach to Refine the Inhalation Risk Assessment for Point of Contact Toxicity: A Case Study Using a New Approach Methodology (NAM) December 4 and 6, 2018 FIFRA Scientific Advisory Panel Meeting". U.S. Environmental Protection Agency). Frohlich, E., Mercuri, A., Wu, S., and Salar-Behzadi, S. (2016). Measurements of Deposition, Lung Surface Area and Lung Fluid for Simulation of Inhaled Compounds. Front. Pharmacol. 7, 181. Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B., and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867. Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The effect of aging on nasal mucociliary clearance, beat frequency, and ultrastructure of respiratory cilia. Am. J. Respir. Crit. Care Med. 163, 983-988. Joki, S., and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea-pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169. National Research Council (1988). Air Pollution, the Automobile, and Public Health. Washington, DC: The National Academies Press. Paul, P., Johnson, P., Ramaswamy, P., Ramadoss, S., Geetha, B., and Subhashini, A. (2013). The effect of ageing on nasal mucociliary clearance in women: a pilot study. ISRN 2013, 598589. Petersen, E.J., Sharma, M., Clippinger, A.J., Gordon, J., Katz, A., Laux, P., et al. (2021). Use of Cause-and-Effect Analysis to Optimize the Reliability of In Vitro Inhalation Toxicity Measurements Using an Air–Liquid Interface. Chem. Res. Toxicol. 34, 1370–1385. Sharma, G., and Goodwin, J. (2006). Effect of aging on respiratory system physiology and immunology. Clin. Interv. Aging 1, 253-260. AOPWiki 2021-09-28T04:51:39

2023-04-29T16:03:05